
























































































































































COPYRIGHT DEPOSIT. 















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/ 






















THE UNIVERSITY OF CHICAGO PRESS 
CHICAGO, ILLINOIS 


THE BAKER & TAYLOR COMPANY 

NEW YORK 

THE CAMBRIDGE UNIVERSITY PRESS 

LONDON 

THE MARUZEN-KABUSHIKI-KAISHA 

TOKYO, OSAKA, KYOTO, FUKUOKA, SENDAI 


THE MISSION BOOK COMPANY 

SHANGHAI 


METHODS IN 

PLANT HISTOLOGY 


By 

CHARLES j; CHAMBERLAIN, PhD., Sc.D. 

Professor of Morphology and Cytology in the University of Chicago 


FOURTH REVISED EDITION 



THE UNIVERSITY OF CHICAGO PRESS 
CHICAGO • ILLINOIS 













&KG73 


C4-4 

I'TM. 


Copyright 1901,1905,1915, and 1924 By 
The University of Chicago 


All Rights Reserved 


Published June 1901 
Second Edition October 1905 
Third Edition May 1915 
Second Impression December 1916 
Third Impression July 1920 
Fourth Impression January 1923 
Fourth Edition October 1924 


Composed and Printed By 
The University of Chicago Press 
Chicago, Illinois. U.S.A. 



©Cl A808857 


VO [ 




PREFACE TO THE FIRST EDITION 


This book has grown out of a course in histological technic 
conducted by the author at the University of Chicago. The course 
has also been taken by non-resident students through the Extension 
Division of the University. The Methods were published over a 
year ago as a series of articles in the Journal of Applied Microscopy , 
and have called out numerous letters of commendation, criticism, 
suggestion, and inquiry. The work has been thoroughly revised and 
enlarged by about one-half. It is hoped that the criticism and 
suggestion, and also the experience gained by contact with both 
resident and non-resident students, have made the directions so 
definite that they may be followed, not only by those who work 
in a class under the supervision of an instructor, but also by those 
who must work in their own homes without any such assistance. 

More space has been devoted to the paraffin method than to 
any other, because it has been proved to be better adapted to the 
needs of the botanist. The celloidin method, the glycerin method, 
and freehand sectioning are also described, and their advantages and 
disadvantages are pointed out. 

The first part of the book deals with the principles of fixing 
and staining, and the various other processes of microtechnic, while 
in the later chapters these principles are applied to specific cases. 
This occasions some repetition, but the mere presentation of general 
principles will not enable the beginner to make good mounts. 

The illustrations and notes in the later chapters are not intended 
to afford a study of general morphology, but they merely indicate 
to students with a limited knowledge of plant structures the principal 
features which the preparations should show. The photomicro¬ 
graphs were made from the author’s preparations by Dr. W. H. 
Knap, and Figures 52, 57, and 59 (Figs. 61, 66, and 68 of second 
edition) were drawn by Miss Eleanor Tarrant; all other figures of 
plant structures were made from the author’s drawings. 

Corrections and suggestions will be heartily appreciated. 

~ Charles J. Chamberlain 

Chicago 

June 1, 1901 


V 


PREFACE TO THE SECOND EDITION 


It is gratifying to the author to learn that the kindly reception 
accorded to Methods in Plant Histology has exhausted the edition. 
Since the first edition appeared, a little more than four years ago, 
laboratory methods have been greatly improved, and systematic 
experiments have* made it possible to give much more definite 
directions for making preparations. 

In the present edition much more attention has been given to 
collecting material. Professor Kleb’s methods for securing various 
reproductive phases in the algae and fungi have been outlined in a 
practical way. Methods for growing other laboratory material are 
more complete than in the earlier edition. 

The paraffin method has been much improved, and the glycerin 
method has been almost entirely replaced by the Venetian turpentine 
method, to which a whole chapter is devoted. Other new chapters 
deal with michrochemical tests, freehand sections, special methods, 
and the use of the microscope. 

The author is deeply indebted to his colleague, Dr. W. J. G. 
Land, for numerous suggestions and improvements in methods. 

Corrections and suggestions will be heartily appreciated. 


Chicago 
July 1, 1905 


Charles J. Chamberlain 


VI 


PREFACE TO THE THIRD EDITION 


The continued appreciation accorded to Methods in Plant His - 
tology has exhausted the second edition. Since that edition appeared, 
methods have become more and more exact, so that the present 
volume is practically a new book. The general arrangement of the 
subject-matter, and directions for collecting material and for secur¬ 
ing reproductive phases in the algae and fungi have been retained, 
and a chapter on “Photomicrographs and Lantern Slides” (chap, xii) 
has been added. 

Great improvements have been made in the paraffin method, so 
that sections are easily cut which were impossible ten years ago, 
while ten years of added experience with the Venetian turpentine 
method have made it possible to describe it so definitely that even 
the beginner should find no serious difficulty. 

The author is deeply indebted to his colleague, Dr. W. J. G. 
Land, for numerous suggestions and improvements covering the 
whole field of microtechnic. He is also greatly indebted to Dr. S. 
Yamanouchi for many improvements in the methods applicable to 
algae and mitotic figures. 

Corrections and suggestions will be heartily appreciated. 

Charles J. Chamberlain 

Chicago 

May, 1915 


PREFACE TO THE FOURTH EDITION 

It is gratifying to the author that the appreciation accorded to 
Methods in Plant Histology , when it first appeared as a series of 
articles in the Journal of Applied Microscopy , has continued through 
three editions of the book. While the chapter headings and general 
arrangement remain about the same as before, the book has been 
almost entirely re-written. 

Directions for collecting material have been amplified and the 
preparation of the most familiar laboratory types has received 
particular attention. While no radical changes have been made in 
the paraffin method, the process has been shortened and improved; 
the Venetian turpentine method, introduced in the second edition 
and improved in the third, has come into such general use that the expe¬ 
rience of many laboratories has been added to that of our own, and the 
directions have become so definite that there is little excuse for fail¬ 
ures. The cellulose acetate method, which may do as much for woody 
structures as the Venetian turpentine method has done for its class of 
mounts, is outlined in a tentative way, and the chapter on “ Photo¬ 
micrographs and Lantern Slides” has been extended and improved. 

The introduction of American stains, which are becoming very ac¬ 
curately standardized, has occasioned some modifications throughout. 

The author is even more deeply indebted than before to his 
colleague, Dr. W. J. G. Land, for suggestions and improvements 
covering the whole field of microtechnic and photography. He is 
also indebted to Dr. S. Yamanouchi for improvements applicable to 
algae and mitotic figures. Dr. Paul J. Sedgwick is responsible for 
much of the improvement in photomicrography and for many of 
the photomicrographic illustrations. To Miss Ethel Thomas, who 
assisted me for many years, I am indebted for improvements, criti¬ 
cisms, and suggestions covering the whole range of the book. Besides, 
I must thank a host of colleagues and students all over the world for 
help in all phases of the subject. 

Corrections and suggestions will be heartily appreciated. 

Charles J. Chamberlain 


viii 


Chicago 
October, 1924 


CONTENTS 

PART I 

PAGE 

Introduction. 3 

Chapter I. Apparatus. 5 

Chapter II. Reagents.18 

Killing and Fixing Agents.18 

Dehydrating Agents.33 

Formulas for Alcohols.35 

Clearing Agents.36 

Miscellaneous Reagents.39 

Chapter III. Stains and Staining.41 

The Haematoxylins.43 

The Carmines.52 

The Anilins.54 

Combination Stains.63 

Chapter IV. General Remarks on Staining.69 

Selection of a Stain.69 

Theories of Staining.70 

Practical Hints on Staining.73 

Chapter V. Temporary Mounts and Microchemical Tests . . 76 

Chapter VI. Freehand Sections.83 

Chapter VII. The Glycerin Method.96 

Chapter VIII. The Venetian Turpentine Method .... 101 

Chapter IX. The Paraffin Method.107 

Killing and Fixing.107 

Washing.109 

Hardening and Dehydrating.110 

Clearing.Ill 

The Transfer from Clearing Agent to Paraffin.112 

The Paraffin Bath.113 

Imbedding.114 

Cutting.116 

Fixing Sections to the Slide.118 

Removal of the Paraffin.120 

ix 
































X 


CONTENTS 


PAGE 

Removal of Xylol or Turpentine.121 

Transfer to the Stain.121 

Dehydrating. .121 

Clearing.122 

Mounting in Balsam.122 

A Tentative Schedule for Paraffin Sections.123 

Chapter X. The Celloidin Method .124 

Chapter XI. The Cellulose Acetate Method .130 

Chapter XII. Special Methods .132 

Very Large Sections.132 

Stony Tissues.133 

Petrifactions.133 

Thick Sections.135 

Land’s Gelatin Method.137 

Schultze’s Maceration Method.137 

Jeffrey’s Maceration Method.138 

Protoplasmic Connections.138 

Staining Cilia.141 

Chondriosomes.142 

Canaliculi.143 

Vascular Bundles in Living Tissues.144 

Staining Living Structures.144 

Chapter XIII. Photomicrographs and Lantern Slides . . * . 145 

Photomicrographs.145 

Lantern Slides.152 


PART II 

Specific Directions.163 

Chapter XIV. Myxomycetes and Schizophytes. 164 

Chapter XV. Chlorophyceae.173 

Chapter XVI. Phaeophyceae.198 

Chapter XVII. Rhodophyceae. 205 

Chapter XVIII. Fungi. 209 

Phycomycetes.209 

Hemiascomycetes.215 

Ascomycetes.216 

Lichens.221 

Basidiomycetes.221 



































CONTENTS 


xi 


PAGE 

Chapter XIX. Bryophytes—Hepaticae.229 

Chapter XX. Bryophytes— Musci .238 

Chapter XXI. Pteridophytes—Lycopodiales.244 

Chapter XXII. Pteridophytes—Equisetales.251 

Chapter XXIII. Pteridophytes—Filicales.254 

Chapter XXIV. Spermatophytes—Gymnosperms.269 

Cycadales.269 

Ginkgoales.276 

Coniferales.278 

Gnetales.289 

Chapter XXV. Spermatophytes—Angiosperms.290 

Chapter XXVI. Using the Microscope.312 

Chapter XXVII. Labeling and Cataloguing Preparations . . 318 

Chapter XXVIII. A Class List of Preparations.320 

Chapter XXIX. Formulas for Reagents.327 

Bibliography.340 

Index.347 




















PART I 



























INTRODUCTION 


The pollen grain of a lily, placed on a dark background, is barely 
visible to the naked eye; but with modern technic, such a pollen 
grain can be cut into fifty sections, the sections can be mounted and 
stained without getting them out of order, a photomicrograph can 
be made from the preparation and a lantern slide from the photo¬ 
micrograph, and finally there appears upon the screen a pollen grain 
10 feet long, with nuclei a foot in diameter, nucleoli like baseballs, 
and starch grains as large as walnuts. With such preparations, 
botanists are now showing clearly the nature of structures which, 
only a few years ago, were good subjects for philosophical speculation. 

To become a good technician, the student should follow carefully 
every detail of the various schedules and, when the routine becomes 
familiar, he should try to understand the reason for every step. In 
some cases, a dozen different schedules might have been given. 
When fundamentals have been grasped, the student will be able to 
make such variations as individual cases may require. 

Those who regard the making of mounts as mere mechanical 
drudgery which can be done by an assistant are likely to become 
armchair investigators, drawing false conclusions or becoming 
scholastic grafters, according as the assistant is mediocre or talented. 
Besides, there is always the danger that a talented but underpaid 
assistant may “hold out” something. Some time-honored theories 
would have been abandoned long ago if certain prominent investi¬ 
gators had not relied upon comparatively untrained assistants for 
their mounts. Benjamin Franklin’s advice, “If you would have 
your business done, go; if not, send,” applies very well to the case 
in hand. 

We strongly advise the student to collect his own material in the 
field, for such collecting is a valuable part of a botanical education. 
There are details of habitat and behavior which are never described 
in books. One learns gradually, by experience, that certain kinds 
of plants grow in certain kinds of places; and further, that not only 
the season, but even the weather may be an important factor. A 
heavy rain may cause some algae to disappear; while the same rain, 

3 


4 


INTRODUCTION 


followed by a few days of sunshine, will bring ideal conditions for 
collecting Myxomycetes and many other fungi. One learns that 
while Lycogala is pink, it is in the free nuclear condition, and that 
Stemonitis is in that condition as long as it is white; that Volvox may 
be abundant at the bottom of a pond when there is scarcely any in 
suspension. The successful investigator learns how a flower bud 
should look, if it is to yield floral development, how the flower looks 
when the embryo sac is mature; and how it looks after fertilization. 

Such studies add immensely to the value of a preparation. A 
considerable part of a botanical education can be gained by collecting 
material, making and studying preparations, reading what is avail¬ 
able, and thinking. 

Finally, do not imagine that you must always make an elaborate 
permanent mount before you examine anything with the microscope. 
Look at living material whenever possible; make freehand sections or 
tease with needles, and thus make that preliminary survey which 
should always precede the study of permanent mounts. 


CHAPTER I 


APPARATUS 

The amount of apparatus required for histological work varies, 
temporary mounts, glycerin mounts, and freehand sections requiring 
only a razor and a microscope, while the paraffin method, which 
represents the highest development of technic, brings into use nearly 
all the equipment of the histological laboratory. The following list 
includes only the apparatus necessary for making preparations: 
a microscope; a microtome; a razor; a hone and a good razor strop; 
a paraffin bath; a turntable; a scalpel; a pair of needles; a pair of 
scissors; a pair of forceps; staining-dishes; solid watch glasses; 
bottles; a graduate (50 or 100 c.c.); pipettes; slides, 1X3 inches; 
round covers, 18 mm. or f inch in diameter; and square covers, £ inch. 
Longer covers will be needed for some of the serial sections. 

A microscope should have a rack and pinion coarse adjustment, 
a fine adjustment, two eyepieces magnifying about four and eight 
diameters, a low-power objective of about 16-mm. focus, and a 
high-power objective of about 4-mm. focus, a double nosepiece, an 
iris diaphragm, and an Abbe condenser. A cheap and practical 
form is shown in Figure 1, and similar instruments are for sale by all 
the leading companies. 

Since the chemicals used in histological technic are likely to 
damage the stage and substage of the microscope, it is well to place 
upon the stage a piece of glass 3 or 4 inches square. A lantern- 
slide cover is just right for this purpose. It is not necessary to fasten 
it to the stage, since it is merely for protection while examin¬ 
ing slides which are wet with reagents. In our own laboratory we 
use for examining wet slides a cheap microscope with only a single 
low-power objective and a single ocular. 

Some knowledge of the structure and optics of the microscope is 
necessary if one is to use it effectively. Why are there so many 
diaphragms ? Why is there an arrangement for raising and lowering 
the condenser ? Why does the mirror bar swing ? Why is one side 
of the mirror plane and the other concave? Everyone who uses 


5 


6 


METHODS IN PLANT HISTOLOGY 



even a cheap microscope should know the answers to questions like 
these. All the leading manufacturers furnish, free of charge, booklets, 
explaining the construction of the microscope and giving practical 

directions for its care and use. 

Aside from the microscope 
itself, the microtome is the 
most important piece of ap¬ 
paratus in the laboratory. In 
recent years there has been 
considerable improvement in 
microtomes, but we still have 
two general types—the sliding 
and the rotary. 

The cheapest microtomes 
which have proved to be 
efficient for general work are 
simple forms of the sliding 
microtome like the one shown 


in Figure 2. It should be 
provided with a clamp which 
will hold any kind of a knife 
(Fig. 3). For large or hard 
objects the weakness of these 
small instruments is evident 
from the figures, but such an 
instrument is very useful; it 
is small and is easily carried 
around; it does not get out of 
order; it cuts celloidin well 
and is good for wood sections, 
except the hardest; and it will 
do for general paraffin work. 

Where expense is not too great an objection, a larger microtome 
should be secured. There is great difference of opinion as to the 
relative merits of the sliding and rotary types. As far as convenience 
and rapidity are concerned, the rotary microtome is unquestionably 
superior; further, it will produce good sections with less care and skill, 
because the movements are automatic. The fact that a ribbon carrier 
is so easily used with the rotary is another great advantage. But 


Fig. 1.—An efficient microscope of moderate price. 
The leading optical companies put the same objectives 
and oculars upon such instruments as upon their most 
expensive stands. 


APPARATUS 


7 


the sliding microtome also has its advantages. Obviously, for sec¬ 
tions of stems and general celloidin work, where the knife is used in a 



Fig. 2.—The student’s microtome 




very oblique position, 
it is not only superior, 
but it is the only type 
which has proved to be 
efficient. Attempts to 
place the knife in an 
oblique position in ro¬ 
tary microtomes have 
not been encouraging. 

For very thin paraffin 
sections the advan¬ 
tages of the sliding 
microtome are such 
as appeal only to the 
expert. With both ro¬ 
tary and sliding types, 
a little of the paraffin 
is sure to stick to the side of the knife next the object after every 
section. Unless this be wiped off, the face of the block is dragged 



Fig. 3.—Clamp to hold an ordinary razor 
or a heavy microtome knife. 







8 


METHODS IN PLANT HISTOLOGY 


across it and the next section is damaged even before it is cut. The 
side of the knife next the object should be wiped with the finger, 
theoretically after every section. It is very inconvenient to wipe 
the knife in a rotary microtome. Another advantage of the sliding 
type is easy to feel but difficult to describe: In the rotary micro¬ 
tome the stroke is so automatic that there is little room for skill, but 
in the sliding microtome, with one’s hand on the sliding block, little 
variations in the stroke, variations which become instinctive, give the 
expert a control not yet attained in the rotary forms. 

Amateurs, and even professional botanists who have little 
aptitude in the use of machines, had better rely upon the rotary 
microtome. However, no better comment on the comparative merits 
of the two forms could be given than the practice of an expert tech¬ 
nician in our own laboratory, who uses a rotary microtome when 
making sections for ordinary class work, but who turns to a sliding 
microtome of the Jung-Thoma pattern when cutting sections for 
his own research. 

A much-improved rotary microtome, devised by Dr. H. N. Ott, 
president of the Spencer Lens Company, combines the precision and 
stability of the sliding microtome with the convenience of the rotary. 
Its effectiveness depends, in large measure, upon the simple but rigid 
clamp for holding the object. The same firm has also produced a 
very rigid sliding microtome, embodying features suggested by Pro¬ 
fessor It. B. Thomson, of the University of Toronto. The stability, 
which makes such a microtome desirable for cutting sections of hard, 
woody structures, is an advantage in cutting very thin sections of 
material imbedded in paraffin. 

Microtome knives are available everywhere and, when perfectly 
sharpened, are unsurpassed. Those who sharpen knives for sur¬ 
geons can grind out nicks, but they do not know how to sharpen a 
microtome knife and they cannot be taught; they sharpen knives for 
Dr. Carver and Dr. Cutterout. 

In recent years several clamps have been devised to hold the 
blade of the Gillette safety razor, the hard, even edge of which is 
very satisfactory for microtome sections. After dealers had ignored 
our suggestions, Mr. A. W. Strickler, at our request, devised the form 
of holder 1 shown in Figure 4. It is made of brass and can be used in 
either rotary or sliding microtomes. The sectional view shows that 

1 At 1320 East Fifty-seventh Street, Chicago, Illinois. 


APPARATUS 


9 


the two pieces of the holder are curved, a feature which insures great 
rigidity. It is neither necessary nor desirable to have pins fitting 
the three holes in the blade, since they add nothing to the rigidity 
and even interfere with the insertion and adjustment of the knife. 
The knife should not project more than a millimeter beyond the 
holder. The clamp for holding Gillette blades in the Spencer rotary 



Fig. 4. —Strickler’s clamp for holding Gillette blades in the Bausch and Lomb rotary microtome 


microtome is necessarily much heavier (Fig. 5). With the Gillette 
blade in these holders we have cut smooth sections, 2 and 3 i± in thick¬ 
ness, and have cut large sections 2 cm. in diameter and 15 m in thick¬ 
ness, even such refractory objects as the strobili of Isoetes and 
Selaginella cutting as smoothly as with a first-class microtome knife. 



Fig. 5. —Clamp for holding Gillette blades in the Spencer rotary microtome 

When the success of the holder—or rather, its sale—became evident, 
two prominent optical companies, without any apologies or reference 
to Mr. Strickler, began to manufacture it and advertised it in their 
catalogues. Their holders are inferior to Mr. Strickler’s, doubtless 
because they overlooked a very important, but very obscure, detail. 

Mr. Ralph B. Larsen, 7126 Woodlawn Avenue, Chicago, Illinois, 
makes these holders, selling the holder for the Bausch and Lomb 
rotary microtome at $5.00, and the heavier holder for the Spencer 
rotary microtome at $7.50. 

Many have trouble with the holders, even as made by Mr. Larsen, 
because they fail to remember that the blade is not straight, like a 
microtome knife, but is bent into a curve. Consequently, the 


















10 METHODS IN PLANT HISTOLOGY 

holder should not be inclined toward the paraffin as far as the micro¬ 
tome knife is inclined. The exact angle cannot be given, since it will 
vary with the amount the blade is allowed to project beyond the holder. 
But, just as in case of the microtome knife, which, when sharpened 
with a back as it always should be, has a double 
bevel like that of the safety-razor blades, the angle 
should be as near the vertical as possible. With 
either a microtome knife or blade in a holder, the 
paraffin block will hit the shoulder of the bevel after 
the section is cut if the position be too nearly 
vertical. A study of Figure 6 should enable anyone 
to get the proper orientation. 

When the Gillette blade begins to lose a little of 
its effectiveness for microtome work it will make 
two or three scalpels. With a pair of stout shears, 
cut the blade into pieces, as indicated in Figure 7. Take a small 
steel nail and flatten the head and upper part by laying it upon a 
piece of iron and hitting it with a hammer, or by squeezing it in a vise; 



Fig. 6. —Proper 
relative positions of 
holder, Gillette blade, 
and paraffin. 





3 


Fig. 7. —Scalpels made from Gillette blades, showing a blade which has been cut into pieces 
with shears, three of the pieces soldered to nails with flattened heads, and one of the pieces used in 
an ordinary needle-holder. 


then solder the blade to the nail, and use the scalpel in an ordinary 
needle-holder, or drive the nail into any wooden holder. A dozen of 
these scalpels can be made in ten minutes. 







































APPARATUS 


11 


The stout razors our grandfathers used to shave with are excellent 
for freehand sectioning and even for cutting sections on the micro¬ 
tome. The blade should be flat on one side 
(Fig. 8A). Modern razors (Fig. 8 B), with 
delicate blades ground hollow on both sides, 
are worthless for cutting sections of plants. 

There should be two good hones: a 
fine carborundum hone for the preliminary 
sharpening, and a yellow Belgian hone for 
finishing. About 10X2| inches is a good 
size. If the second hone be quite hard 
and the finishing skilfully done, little or 
no stropping may be necessary. The best 
strops used by barbers are satisfactory for microtome knives. 

There are numerous forms of the paraffin bath. Those with a 
water-jacket, a thermometer, and a thermostat to maintain an even 
temperature are the most convenient. 



Fig. 8.—The type shown in A 
is good for microtome work; that 
shown in B is worthless for micro¬ 
tome work, but can be used for 
freehand sections of leaves. 



Fig. 9. —Land’s electric constant apparatus, showing diagram of the automatic switch, as 
described in the Botanical Gazette, November, 1911. 


A bath with a water-jacket and the electric thermostat devised 
by Dr. Land maintains an even temperature, in spite of any variations 
in the current. The appearance and principal features of this 
thermostat are shown in Figures 9 and 10. A detailed description of 



























12 


METHODS IN PLANT HISTOLOGY 


this thermostat is given in the Botanical Gazette for November, 1911. 
One familiar with tools and electricity could make this thermostat 
at a very moderate outlay. Unless the coil in the heater is perfectly 
protected, there will be a short circuit. This danger can be obviated, 
in large measure, by using oil instead of water in the jacket. 



Fig. 10.—Thermostat, heater, and switch of Land’s electrical constant apparatus 


Another form of heater, using long electric bulbs instead of a 
coil, can be made with less skill. From a sheet of transite f inch thick, 
make a box just the size of the bottom of the bath and about 4 inches 
deep. In this box put three long electric bulbs. Two of the bulbs, 
about 250 watt, giving a temperature of about 40° C., should be 
connected with the regular outlets and should be on all the time. 












APPARATUS 


13 


The third bulb, which should be capable of raising the tempera¬ 
ture 15° or 20° C., should be connected with a thermostat. An ef¬ 
ficient thermostat, capable of keeping the variation in temperature 
within about 1° can be made in a short time. Take a piece of 
soft steel f inch wide and T V inch thick, and 17 inches long; lay upon 
it three strips of aluminium of the same width and length, but only 
-fa inch thick. These four pieces are fastened together by drilling 
numerous holes, not more than fa inch in diameter and riveting with 
pieces of brass wire. After fastening the four pieces together for 
about 7 inches, begin to bend all four pieces, fastening them together 
as the bending continues, so that finally there will be a horseshoe 
shape with the parallel sides about 1J inches apart. One end of 



the horseshoe is fastened to a block of transite or other non-conductor, 
while the other end moves freely. On this free end is fastened a good 
platinum contact, which can be bought at any automobile supply 
store. This contact should be on a screw, so that it may be adjusted. 
A similar contact is fastened to a post, so that the two contacts are 
about | inch apart. Anyone familiar with electricity can make the 
connections. Various temperatures are secured by changing the 
distance between the platinum points. To prevent sparking, there 
must be a condenser in the circuit leading to the thermostat. A 
telephone condenser (0.500MF) is satisfactory. Possibly, some of the 
cheap radio condensers would do. 

A bath which, if carefully watched, gives the very best results, 
can be made by any tinner (Fig. 11). It is simply a triangular piece 












14 


METHODS IN PLANT HISTOLOGY 


of brass, 2 or 3 mm. in thickness, with three legs screwed into it. There 
should be long brass boxes to contain the paraffin. It is neither neces¬ 
sary nor desirable that the boxes have covers. Boxes are easily made 
from square brass tubing, 1 inch square. Cut the tube into pieces 
6 inches long, saw off one side, and solder pieces into the ends with 
hard solder. The brass plate can be heated with any kind of a flame 
at the pointed end. 

Since the Venetian turpentine method has almost entirely dis¬ 
placed the glycerin method, the turntable is becoming an unfamiliar 
object in the botanical laboratory; but some objects, like Nemalion 
and moss protonema, are still mounted in glycerin jelly, and so one 
still finds occasional use for this once necessary apparatus. A service¬ 
able form is shown in Figure 12. The more expensive turntables, 



Fig. 12.—Turntable 


with devices for automatic centering, present no practical advantages 
and the centering devices are often in the way. 

Scalpels made from thin razor blades have been mentioned 
already. For trimming paraffin blocks and handling paraffin ribbons 
a more rigid scalpel is necessary. A scalpel with a straight edge is 
preferable. 

Needles are used so constantly that it is well to have clamping 
holders. However, if it were not for the trouble of inserting and 
pulling out needles, nothing is quite equal to a rather large handle 
whittled out from a piece of light pine. 

Scissors are seldom used in the botanical laboratory except for 
cutting out labels. Rather stout scissors, with blades about 2| inches 
in length, are best for general purposes. 

It is convenient to have two pairs of forceps, a strong pair for 
handling slides and a delicate pair for handling covers. If there is 




APPARATUS 


15 


to be only ,one pair, they must be strong enough for the slides but 
not too clumsy for covers. Curved forceps are not necessary; the 
cover-glass forceps, used by bacteriologists in staining on the cover, 
are of no use in botanical work. 

Stender dishes are very generally used in staining on the slide. 
The form shown in Figure 13A, about 60X90 mm., is in quite general 
use. Some prefer the Coplin jar, shown in Figure 135. The latter 
is more troublesome to clean, but requires less of the reagent. Many 
other forms are bn the market. When large numbers of slides of the 
same kind are to be stained at one time, small battery jars, holding 
about a liter, may be used. In this case, it is well to have a rack, 
holding about thirty slides, so that all may be transferred at the 
same time from one reagent to another. With this convenience, it 
is not necessary to handle the slides separately, except at critical 
stages. 



A 


B 


Fig. 13.—Staining-dishes: A, Stender dish; B, Coplin jar 


A very cheap and practical device, which can be used in an 
ordinary Stender dish, is shown in Figure 14. It is simply a coil of 
brass wire, 0.064 inch in diameter (No. 14, Band’s gauge), wound so 
that the coil is about f inch across. Such a coil, carrying 15 slides, 
will go into an ordinary Stender dish, except that the coil projects 
enough to prevent the cover from fitting. Taller glasses, from the 
five and ten cents store, can be used for the absolute alcohol and xylol, 
which must be kept well covered. We have been using a coil made 
from wire 0.051 inch in diameter (No. 16 Band’s gauge), wound so 
that the coil is if inch across. It holds the slides and the ordinary 
Stender dish can be covered. 




























16 


METHODS IN PLANT HISTOLOGY 


Solid watch glasses, or Minots, as they are often called, are always 
useful. Each student should have a dozen or more. 

Each student should have three bottles of about 1-liter capacity 
for 90 per cent alcohol, absolute alcohol, and xylol. In addition, a 
half-dozen bottles, holding about 100 c.c., will be useful. There 
should be two bottles, holding about 50 c.c., for clove oil. If one is 



Fig. 14.—Coil of brass wire holding 15 slides 


doing much research work, it will be convenient to have many more 
bottles for graded series of alcohols and xylols. 

There should be a graduate, preferably 50 c.c. or 100 c.c. If 
the bottles are of uniform sizes, 50 c.c., 100 c.c., 500 c.c., and 1,000 c.c., 
the student should soon be able to estimate with sufficient accuracy 
for making up reagents which do not require extreme accuracy. 

Three or four pipettes, or medicine-droppers, will be useful. 
Occasionally, the glass of an ordinary pipette thrust into a small 
camera bulb will save time in drawing off reagents. 







APPARATUS 


17 


Slides and covers are a constant expense. Many slides now 
upon the market are imperfect. Beware of slides which are not 
perfectly flat. Be skeptical in regard to any claim that slides are 
already clean enough to use. Of course, there should be no bubbles. 
“ White” slides are to be preferred to those which appear greenish in 
the box. For ordinary class work, slides of medium thickness are 
more serviceable, but for critical cytological work many investigators 
prefer very thin slides. 

There is never any objection to very thin covers, except that they 
require care in cleaning. For mounts which are to be used with an 
immersion lens, it is better to have the cover of the same width as 
the slide. The advantage is evident, since there is no danger of 
getting balsam on the cover when wiping off the immersion fluid; 
besides, one can put sections to the very edge of the slide and still 
be sure that they will be covered. Since most mounts for research 
work are mounted under long covers and are intended for examina¬ 
tion with immersion lenses, we should recommend covers of 25X50 
mm., or even 25X60 mm. Round covers are desirable only when 
mounts are to be sealed on a turntable. Larger slides and corre¬ 
spondingly larger covers are needed for special purposes. 

By consulting a catalogue, which will be furnished by any dealer, 
the beginner can determine what he needs to buy, and what he can 
find substitutes for, if it is necessary to be very economical. 


CHAPTER II 

REAGENTS 

No really new reagents have come into general use since the 
third edition of this book was published in 1915, but there have been 
valuable modifications of some of the older formulas, and improve¬ 
ments in the use of time-honored combinations. The following 
account includes those which are used constantly and also a few 
which are used occasionally. The MicrotomisVs Vade-Mecum, by 
Lee, is written from the standpoint of the zoologist, but it contains 
very complete formulas for stains and other reagents, which are just 
as useful to the botanist. 

A list of reagents, with the quantities used by the average student 
in a three months’ course in methods, is given in chapter xxix. 
“Stains and Staining” are described in chapter iii. 

KILLING AND FIXING AGENTS 

Usually the same reagent is used for both killing and fixing. 
The purpose of a killing agent is to bring the life-processes to a sud¬ 
den termination, while a fixing agent is used to fix the cells and their 
contents in as nearly the living condition as possible. The fixing 
consists in so hardening the material that the various elements may 
retain their natural condition during all the processes which are to 
follow. Zoologists often use chloroform or ether for killing an 
organism, and then use various fixing agents for various tissues. 
No promptings of humanity restrain the botanist from the vivisection 
of plants, but separate reagents for killing and fixing are sometimes 
used, e.g., material may be killed by placing it for a short time in 
Flemming’s fluid, which is a very rapid killing agent, after which the 
fixing may be completed in a chromo-acetic solution, without any 
osmic acid, thus securing the advantages of a very rapid killing 
without the blackening which results from a prolonged treatment 
with a solution containing osmic acid. 

Probably no process in microtechnic is in more urgent need of 
improvement than this first step of killing and fixing. Nearly all 

18 


REAGENTS 


19 


of our formulas are merely empirical, for very few botanists are 
expert chemists, and those who have the requisite knowledge of 
chemistry are interested in physiological problems rather than in 
microtechnic. The principal ingredients of the usual killing and 
fixing agents are: alcohol, chloroform, chromic acid, bichromate of 
potash, potassium iodide, copper acetate, acetic acid, osmic acid, 
formic acid, picric acid, sulphuric acid, platinum chloride, iridium 
chloride, corrosive sublimate, and formalin. We shall consider first: 

THE ALCOHOLS 

a) Ninety-five Per Cent Alcohol. — This is in quite general use 
for material which is needed only for rough work. It is extremely 
convenient, since it kills, fixes, and preserves at the same time and 
needs no changing or washing. It really has nothing to recommend 
it for fine work. It causes protoplasm to shrink, but cell walls 
usually retain their position, so that 95 per cent alcohol will do for 
freehand sections of wood and many herbaceous stems, where it is 
not necessary to preserve cell contents; but even freehand sections 
of tender stems, like geraniums and begonias, will look better if 
better reagents are employed. Alcohols weaker than 95 per cent 
are not to be recommended as fixing agents, although 70 per cent al¬ 
cohol, or even 50 per cent, will preserve material for habit work. 
The time required for fixing in 95 per cent alcohol is about the 
same as for absolute alcohol. The subsequent treatment is the same, 
except that material to be imbedded in paraffin or celloidin must 
be dehydrated in absolute alcohol. Material preserved in weaker 
alcohols and intended only for habit study may be kept in the reagent 
until needed for use. Unless some glycerin be added, material left in 
95 per cent alcohol becomes very brittle. Stems, roots, and similar 
objects may be kept indefinitely in a mixture of equal parts of 95 per 
cent alcohol and glycerin. 

Methyl alcohol, or wood alcohol as it is commonly called, serves 
equally well. 

b) Absolute (100 Per Cent) Alcohol.—This is a fair killing and 
fixing agent, it causes but little shrinking of the protoplasm, and is 
a time-saver if material is to be imbedded in paraffin. The time 
required for fixing in alcohol is very short. For small fungi, like 
Eurotium, 1 minute is long enough. Root-tips of the onion, anthers 
of the lily, and similar objects require 15 to 30 minutes. Larger 


20 


METHODS IN PLANT HISTOLOGY 


objects may require an hour. No washing is necessary, but all 
plant tissues contain water; consequently, if material is to be 
imbedded in paraffin, the alcohol used for fixing should be poured 
off and fresh alcohol added before proceeding with the clearing. If 
material is to be mounted in Venetian turpentine, as is likely to be the 
case in small filamentous fungi, the transfer to the stain may be 
made directly from the absolute alcohol to any stain dissolved in an 
alcohol not weaker than 85 per cent. Small forms with no vacuoles 
may be transferred to a weaker alcoholic stain or even to an aqueous 
stain; but neither the fixing nor the rude transfer would be at all 
satisfactory with forms like Zyghema or Saprolegnia. 

Acetic acid is used with alcohols to counteract the tendency to 
shrink. One of the most widely known of the alcohol combinations is 


c) Carnoy’s Fluid.— 

Absolute alcohol. 6 parts 

Chloroform. 3 parts 

Glacial acetic acid. 1 part 


The penetration of the reagent is very rapid. An object like an 
onion root-tip is doubtless killed in less than a minute, and 10 to 15 
minutes is long enough for fixing an object of this size. Wash in 
absolute alcohol, changing frequently, until there is little or no odor 
of acetic acid. For a root-tip, the entire process of fixing and wash¬ 
ing should not require more than an hour. It is better to imbed in 
paraffin at once, but when. this is not convenient the material may 
be washed in abolute alcohol until the odor of acetic acid disappears, 
cleared in xylol and, with a block of paraffin about half the bulk of 
the liquid added, may be left indefinitely. Cyanin and erythrosin, 
fuchsin and iodine green, and similar combinations give particularly 
brilliant staining after this reagent. 

d) Acetic Alcohol.—Farmer and Shove recommend for fixing 
root-tips of Tradescantia virginica a mixture of two parts absolute 
alcohol and one part of glacial acetic acid. The mixture is allowed 
to act for 15 to 20 minutes, after which the acid is washed out with 
absolute .alcohol and the material is imbedded as soon as possible. 

e) Formalin Alcohol.—One of the most satisfactory of the alcohol 
combinations is formalin alcohol. Various proportions are used by 
different workers. Professor Lynds Jones, who first brought this 
combination to my notice, added 2 c.c. of commercial formalin to 





REAGENTS 


21 


100 c.c. of 70 per cent alcohol. We have used a larger proportion of 
formalin, often as much as 6 c.c. to 100 c.c. of 70 per cent alcohol. 
Results which seem equally good have been secured by adding 4 to 
6 c.c. of formalin to 100 c.c. of 50 per cent alcohol. Material in this 
fixing agent may be left until needed for use. 

/) Formalin Acetic Alcohol.—This combination seems even better 
than the preceding. We use about 5 c.c. of glacial acetic acid and 
5 c.c. of commercial formalin to 90 c.c. of 50 per cent or 70 per cent 
alcohol. 

When one is on a long trip, moving frequently from place to place, 
with little opportunity to make the numerous changes which are 
necessary when using the chromic formulas, this is the best fixing 
agent we have found. It will fix and preserve an amount of material 
equal to its own weight, and the material may be left in the solution 
for months. The reagent is good for almost any material, except 
the unicellular and filamentous algae and fungi, which are more 
satisfactory in media containing no alcohol. 

THE CHROMIC-ACID GROUP 

Chromic acid, or solutions with chromic acid as a foundation, are 
the most generally useful killing and fixing agents yet known to the 
botanist. A 1 per cent solution of chromic acid in water gives good 
results, but it is better to use the chromic in connection with other 
ingredients, such as acetic acid, formic acid, osmic acid, etc. Chromic 
acid does not penetrate well, and this is one reason why it is seldom 
used alone. Unfortunately it precipitates some liquid albuminoids 
in the form of filaments and networks, which may be mistaken for 
structural elements. In botanical work, acetic acid is nearly always 
mixed with chromic acid. The pickles of the dinner table show that 
acetic acid is a good preservative, and that it causes little or no 
shrinking. It penetrates rapidly, and is likely to cause swelling 
rather than shrinking, thus counteracting the tendency of chromic 
acid to cause plasmolysis. The swelling is as bad as shrinking. 
If the proportion of acetic acid is too high, material may even 
break up; but 2 per cent, or even 3 per cent, may be used to 
show the topography of an embryo sac of an angiosperm, or the 
free nuclear stage of the endosperm of a gymnosperm; and for fila¬ 
mentous algae, which are to be mounted whole, 3 per cent is very 
effective. 


22 


METHODS IN PLANT HISTOLOGY 


It will be found convenient to have in the laboratory the following 
stock solution of chromo-acetic acid from which various solutions 
can be made as they are needed: 


Chromic-acid crystals. 10 g. 

Glacial acetic acid. lOc.c. 

Water. 1,000 c.c. 


To make a solution containing 0.5 g. of chromic acid and 2 c.c. of 
glacial acetic acid to 100 c.c. of water, add 50 c.c. of water to 50 c.c. 
of the stock solution, and then add to the weakened solution 1.5 c.c. 
of glacial acetic acid. Any desired proportions can be secured in a 
similar way. Weighing the crystals for every new proportion is 
more tedious. The proportions of the various ingredients, for the 
present at least, must be determined by experiment. With favorable 
objects like fern prothallia, Spirogyra, and other things which can 
be watched while the fixing is taking place, suitable proportions are 
rather easily determined, because specimens, after being placed in 
the reagent, may be examined at frequent intervals, and combinations 
which cause plasmolysis may be rejected and different proportions 
tried until satisfactory results are secured. For example, fern pro¬ 
thallia might be placed in the following solution: chromic acid, 2 g.; 
acetic acid, 1 c.c.; and water, 97 c.c. If plasmolysis takes place, 
weaken the chromic or strengthen the acetic, since the chromic 
has a tendency to produce contraction, and the acetic to cause 
swelling. A 2 per cent solution of chromic acid is rather strong and 
a \ per cent is rather weak; for most things, 1 per cent seems to be 
about right. For the fern prothallia, the stock solution, with the 
addition of 2 c.c. of glacial acetic to 100 c.c. of the solution, is satis¬ 
factory for material to be mounted whole, and also for sections. 
A combination may be quite satisfactory for fern prothallia and still 
fail to give good results with Spirogyra, and a combination which 
succeeds very well with Spirogyra may not succeed at all with 
Vaucheria. For very critical work the most favorable proportions 
must be determined for the particular plant under investigation. 
In observing the effect of the fixing one can determine whether 
there is any noticeable plasmolysis or distortion, but whether the 
fixing is thorough can be determined only by noting how the tissues 
endure the subsequent processes. When the effect of the reagent 
cannot be observed directly, it is well to make a freehand section 





REAGENTS 


23 


and thus determine whether plasmolysis takes place. It is not 
safe to judge the action of a fixing agent by the appearance of sections 
cut from material which has been imbedded in paraffin, because 
shrinking of the cell contents often takes place during the transfer 
from absolute alcohol to the clearing agent or during infiltration 
with paraffin, and sometimes even during later processes. When 
there is doubt as to proportions, we should suggest 2 c.c. chromic 
acid, 3 c.c. acetic acid, and 300 c.c. water as a good formula for most 
purposes. 

A large quantity of the fixing agent is required and it cannot be 
used again. The volume of the fixing agent should be at least 25 
times that of the material to be fixed. We use about 50 volumes of 
the fixing agent to one of the material. 

The time required for fixing undoubtedly varies with different 
objects, but even a delicate object, like Spirogyra, which is penetrated 
immediately, should remain in the fixing fluid for 18 to 24 hours. 
Most botanists leave material like onion root-tips and lily ovaries 
in the chromo-acetic acid about 24 hours. Some recommend longer 
periods. Christman, in his work on rusts, left material for three 
days in Flemming’s fluid, a much more vigorous agent than the 
chromo-acetic acid. We have often imbedded material which 
had been in chromo-acetic acid for several days, and it seemed to 
have suffered no injury. It is well known that zoologists allow 
fixing agents like Muller’s fluid and Erlicki’s fluid to act for weeks 
before the material is passed on to the next stage, and it may well be 
questioned whether botanists have not made a mistake in allowing 
the chromic solutions to act for so short a time. More rapid pene¬ 
tration, and consequently more immediate killing, can be secured if 
the reagent is kept warm (30° to 40° C.). The warming also shortens 
the time required for fixing, but, for cytological work, it is quite 
possible that the danger of producing artifacts may be increased by 
the heat. 

After fixing is complete, all reagents containing chromic acid 
as an ingredient should be washed out with water. Running water 
is desirable, and where this is not convenient the water must be 
changed frequently. 

About 8 or 10 hours should be long enough for filamentous algae 
and fungi, which are immediately penetrated by the water. It is a 
good plan to start the washing in the morning and let the material 


24 


METHODS IN PLANT HISTOLOGY 


wash all day. For fern prothallia, onion roob-tips, lily anthers, and 
any material from such a size up to cubes a centimeter square, let the 
material wash for 24 hours. Even for delicate algae, 24 hours does 
no damage, and some of the best cytologists prefer the prolonged 
washing. 

Many methods have been devised for insuring thorough washing 
and for facilitating the process. The most obvious method is to 
allow a gentle stream of water to flow into the Stender dish or bottle 
containing the material. There is little danger in this method if the 
material is heavy enough to remain at the bottom: the only objection 
is that much of the water never reaches the bottom where it is needed. 
If material is lighter, tie a piece of cheese-cloth over the mouth of the 
bottle. 

A method devised by Dr. Dudgeon is simple and very efficient, 
especially for delicate material or objects which have a tendency to 
float. A glass tube, 1 inch in diameter, is cut into pieces 2f inches 
in length. The glass is then heated to round off the sharp edges and, 
while the glass is still very hot, one end is flared a little, so that a 
piece of cloth can be tied over or fastened over it with a rubber band. 
Bolting cloth or bolting silk is best because water passes through it 
so readily. A number of these tubes can be placed in a dish not quite 
so deep as the tubes and water can flow into the dish without any 
danger of making the tubes overflow. With material which does not 
settle readily, this method is a great time-saver. 

An apparatus for washing several collections at one time may be 
made as follows: Get a piece of f-inch lead pipe, bore holes about 
or f inch in diameter and about 1| inches apart, put a short rubber 
tube in each hole and the glass part of a pipette in the end of each 
rubber tube. Connect the lead tube with the faucet by a large 
rubber tube. A still better way is to bore T \-inch holes in the lead 
tube, screw into these holes short brass tubes, and then fasten the 
pipettes to the brass tubes with thin rubber tubes. 

If there are no facilities for working with metal, take a wooden 
box about 6 inches wide, 18 inches long, and 4 inches deep; bore 
f-inch holes in the bottom, and into each hole put a piece of rubber 
tubing about 4 or 5 inches in length. The pipettes can be fastened 
in the ends of these rubber tubes. Place the box under the tap. In 
the botanical laboratory at Woods Hole, Massachusetts, large 
quantities of material are washed at one time by using an ordinary 


REAGENTS 


25 


washtub with the bottom arranged as just described for the box. 
If one is using such a large box or tub and does not need all the 
streams of water, the tubes not in use may be closed by means of 
clamps. 

If running water is not available, put the material into a rather 
large bottle or dish; a 200 c.c. bottle is not too large for half a dozen 
J-inch cubes. Change frequently, especially at first. Nothing is 
safe with less than 24 hours of this sort of washing. Where running 
water is not available, Dudgeon’s method is particularly valuable, 
for the tubes can be lifted quickly from one dish into another. 

If the washing has not been thorough, the subsequent staining 
is likely to be unsatisfactory. 

Some of the chromic-acid formulas are as follows: 

a) Stock Chromo-Acetic Solution.— 


Chromic acid. 1 g. 

Glacial acetic acid. 1 c.c. 

Water. 100 c.c. 


This solution has been used quite extensively in embryological 
work upon the higher plants. It fixes thoroughly, but often causes 
plasmolysis in cells with large vacuoles. 

b) Weak Chromo-Acetic Solution (Shaffner’s formula).— 


Chromic acid. 0.3 g. 

Acetic acid. 0.7 g. 

Water. 99.0 c.c. 


This has also been used in embryological work. It causes little 
or no plasmolysis. Difficult material, like Aster heads and ripe 
Capsella pods, cuts more readily after this reagent than after the 
stronger solutions. 

c) Strong Chromo-Acetic Solution.— 


Chromic acid. 1 g- 

Glacial acetic acid. 3 c.c. 

Water. 100 c - c * 


For fern prothallia, most liverworts, moss capsules before they 
have begun to get reddish or brownish, and most filamentous algae 
and fungi, this is a good fixing agent. 











26 


METHODS IN PLANT HISTOLOGY 


d) Special Chromo-Acetic-Osmic Solution.— 


Chromic acid. 1 g. 

Glacial acetic acid. 3 c.c. 

1 per cent osmic acid. 1 c.c. 


We have been trying these three ingredients, in various propor¬ 
tions, for many years and have found this combination very good for 
filamentous algae and fungi, even difficult forms like Hydrodictyon, 
Vaucheria, and Saprolegnia showing practically no plasmolvsis. 
The acetic acid may be too strong for some root-tips and probably 
other objects; but the reagent is worth a trial. With 2 c.c. of osmic 
acid, this seems to be as good for mitosis as Flemming’s solutions, 
and it causes less plasmolysis. Besides, in most cases, no bleaching 


is necessary. 

e ) Licent’s Formula.— 

1 per cent chromic acid. 80 c.c. 

Glacial acetic acid. 5c.c. 

Formalin.. 15 c.c. 


This formula has been recommended for coenocytic algae and 
fungi and for embryo sacs. 

/) Flemming’s Fluid (stronger solution).— 


fl per cent chromic acid. 45 c.c. 

[Glacial acetic acid. 3c.c. 

B. 2 per cent osmic acid. 12 c.c. 


Keep the mixture A made up, and add B as the reagent is needed 
for use, since it does not keep well. This fluid is quite expensive on 
account of the osmic acid. For cytological work it has been very 
popular, and it is especially recommended for chromosomes, centro- 
somes, achromatic structures, and mitotic phenomena in general. 
The fluid should be allowed to act for 24 to 48 hours and the washing 
in water must be very thorough. 

Material should be in very small pieces | inch square, or in thin 
slices | inch or less in thickness, for the fluid penetrates poorly. The 
blackening due to the osmic acid may be removed by peroxide of 
hydrogen just before the slide is passed from the alcohol into the 
stain. Harper and Holden, in their work on Coleosporium, recom¬ 
mended 4 hours on the slide in a 3 per cent solution of the peroxide 
of hydrogen. Some prefer a stronger solution of the peroxide of 











REAGENTS 


27 


hydrogen, even 20 per cent. The peroxide should be in water, if one is 
following it by an aqueous stain, but may be in 50 per cent alcohob 
if it is to be followed by an alcoholic stain. Yamanouchi has used 
chlorine for bleaching, and the results are fully equal to those obtained 
with peroxide of hydrogen, and the chlorine is cheaper. Make the 
bleacher as follows: Place some potassium chlorate crystals—a 
group about as large as a grain of wheat—in the bottom of a 100 c.c. 
Stender dish; add one drop of 25 per cent hydrochloric acid in water; 
immediately fill the Stender full of 30 per cent alcohol and thus 
dissolve the fumes in alcohol. This will bleach sections in 10 minutes, 
or even less. Wash in 30 per cent alcohol 2 or 3 hours before stain¬ 
ing. Trondle uses 1 per cent chromic acid in water for bleaching; 
it is slow, requiring about 8 hours, but he maintains that material 
stains better than after bleaching with peroxide of hydrogen. Accord¬ 
ing to Miss Merriman, the linin in the nuclei of onion root-tips is 
not so well preserved in this solution, but the arrangement of the 
chromatin granules is brought out with greater distinctness. 
Flemming’s safranin, gentian-violet, orange combination gives excel¬ 
lent results after this reagent. 

g) Flemming’s Fluid (weaker solution) .— 


{ 1 per cent chromic acid. 25 c.c. 

1 per cent acetic acid.lOc.c. 

Water. 55 c.c. 

B. 1 per cent osmic acid. 10 c.c. 


As in case of the stronger solution, mix A and B only as needed for 
immediate use. 

Many prefer the weaker solution, because the blackening is not 
so extreme and material does not become quite so brittle. Some 
allow the solution to act for an hour and then transfer the material to 
solution A for about 24 hours. This secures the rapid killing, 
which is the principal virtue of the osmic acid, and avoids the dis¬ 
agreeable blackening, so that little or no bleaching may be necessary. 


h) Benda’s Fluid.— 

1 per cent chromic acid. 16c.c. 

2 per cent osmic acid. 4 c.c. 

Glacial acetic acid. 2 drops 


This modification of Flemming’s stronger solution has been used 
in various investigations upon chromatin. 









28 


METHODS IN PLANT HISTOLOGY 


i ) Merkel’s Fluid.— 

Equal volumes of a 1.4 per cent solution of chromic acid and a 
1.4 per cent solution of platinic chloride. This is also an expensive 
reagent. It is recommended for mitotic phenomena, but does not 


seem to equal Flemming’s solution. 

j) Hermann’s Fluid.— 

1 per cent platinic chloride. 15 parts 

Glacial acetic acid. 1 part 

2 per cent osmic acid. 4 or 2 parts 


This is the most expensive fixing agent yet discovered, and for 
botanical purposes it does not seem to be any better than the cheaper 
chromic mixtures. It is mentioned here with chromic mixtures 
because it originated as a variation of Flemming’s fluid, the platinic 
chloride being substituted for the chromic acid. Recently, it has 
been resurrected and highly recommended for the structure of the 
chromosome. Personally, I do not believe it is equal to Flemming’s 
weaker solution; and, even in this weaker solution, the percentage 
of osmic acid may be too high. The chromo-acetic-osmic solution 
given under (< d ) fixes chromatin very well and, with 2 c.c. of osmic 
acid, seems to be equal to any of the modifications of Flemming’s 
solutions. 

PICRIC ACID 

Use a saturated solution in water or 70 per cent alcohol. One 
gram of picric acid crystals will saturate about 75 c.c. of water or 
alcohol. This reagent penetrates well and does not make the material 
brittle. It is to be recommended when difficulty is anticipated in 
the cutting. If used cold, the time varies from 1 to 24 hours, depend¬ 
ing upon the character of the tissue and size of the specimen. If 
used hot (85° C.), 5 or 10 minutes will be sufficient. Material 
should be washed in 70 or 50 per cent alcohol. Water is injurious, 
and some even go so far as to avoid aqueous stains, unless the ma¬ 
terial has been thoroughly washed. The washing should be continued 
until the material appears whitish and the alcohol no longer becomes 
tinged with yellow. Picro-carmine gives its best results after this 
reagent. Picric acid can be combined with various other fixing agents, 
and so we have picro-sulphuric acid, picro-nitric acid, picro-chromic 
acid, picro-chromic-sulphuric acid, picro-osmic acid, picro-alcohol, 
and picro-corrosive sublimate. The picric acid in all mixtures should 
be rather strong. 





REAGENTS 


29 


A picric-acid combination which has gained some popularity 


for cytological work is 

Bouin’s Fluid.— 

Formalin (commercial). 25 c.c. 

Picric acid (saturated solution in water). 75 c.c. 

Glacial acetic acid. 5 c.c. 


Fix about 24 hours. Rinse in water for a few minutes to remove 
the more superficial picric acid, and then complete the washing in 
35 per cent or 50 per cent alcohol. There is likely to be some swelling, 
but spindles of mitotic figures stain well. The formula has given 
good results with early stages in the female gametophyte of Pinus 
and would be worth a trial with the embryo sacs of angiosperms. 

CORROSIVE SUBLIMATE 

Corrosive sublimate, or bichloride of mercury, is soluble in water 
and in alcohol. About 5 g. will make a saturated solution in 100 c.c. 
of water. It is somewhat more soluble in alcohol, but for practical 
purposes 5 g. in 100 c.c. of 50 per cent alcohol may be regarded as a 
saturated solution. Corrosive sublimate used alone does not give 
as good results as when mixed with acetic acid, chloroform, or picric 
acid. Fixing is very rapid, the material being fixed almost as soon 
as it is penetrated by the fluid. Material which is at all transparent, 
like some ovules and the endosperm of gymnosperms before the 
formation of starch, becomes opaque as soon as fixed, and so the time 
needed for fixing is easily determined. From 10 minutes to one 
hour should be sufficient for onion root-tips or lily ovaries. Smaller 
or larger objects require shorter or longer periods. When used hot 
(85° C.), the fixing is much more rapid. While a few minutes’ 
fixing may be sufficient, we let the reagent reach the boiling-point, 
then remove the flame and just as soon as the bubbling ceases, put 
the material in and leave it until the liquid becomes cool. It may 
be left for hours, without any damage. 

Wash out aqueous solutions with water and alcoholic solutions 
with alcohol. In either case, the washing must be very thorough, 
since preparations from incompletely washed material are sure to be 
disfigured by crystals of corrosive sublimate. After material fixed 
in the aqueous solution has been washed in water for an hour, add 
a little of the iodine solution used in testing for starch. The liquid will 
turn brownish or amber-colored, and then clear up; add a little more, 





30 


METHODS IN PLANT HISTOLOGY 


until the liquid fails to clear up completely, a very slight amber 
remaining for an hour or even permanently. After material fixed 
in the alcoholic solution has been washed in 50 per cent alcohol for 
an hour or more, add the iodine solution used in testing for starch 
or even add, drop by drop, tincture of iodine, until the color fails 
to disappear. With unicellular forms, filamentous forms, and thin 
things, like fern prothallia, such washing is likely to be sufficient, 
but with more bulky material, which is to be sectioned, the crystals 
may appear in the paraffin ribbon. In such cases, the slide should be 
dipped for a minute in the iodine solution just before staining. 

Camphor may be used instead of iodine to hasten the washing, but 
it does not give any color reaction. 

Material should be imbedded as soon as possible, since it gets 
brittle if allowed to remain in alcohol. 

Kinoplasmic ^structures do not stain well with gentian-violet, 
but safranin and the haematoxylins stain almost as well as after 
chromic-acid mixtures, and the carmines give their most brilliant 
stains, as a result of the formation of mercuric carminate. 

The following formulas are merely suggestive: 

a) Corrosive Sublimate and Acetic Acid.— 


Corrosive sublimate. 3 g. 

Glacial acetic acid. 5 c.c. 

Alcohol (50 per cent) or water. 100 c.c. 


b) Corrosive Sublimate, Acetic Acid, and Formalin.— 


Corrosive sublimate . 4 g. 

Glacial acetic acid. 5 c.c. 

Formalin. 5c.c. 

Alcohol (50 per cent) or water. 100 c.c. 


This is our favorite formula. For material which is to be 
mounted in glycerin, glycerin jelly, or Venetian turpentine, use the 
aqueous solution; for material which is to be imbedded, use the 
alcoholic. Cilia are caught and preserved; and even delicate 
organisms, like Volvox, do not collapse. 

c) Corrosive Sublimate, Acetic Acid, and Picric Acid.— 

Corrosive sublimate . 5 g. 

Glacial acetic acid. 5 c.c. 

Picric acid, saturated solution in 50 per cent 

alcohol. 100 c.c. 












REAGENTS 


31 


Miss Ethel Thomas recommends this formula for the female 
gametophyte of Pinus. 

d) Corrosive Sublimate and Picric Acid (Jeffrey’s solution).— 
Corrosive sublimate, saturated solution in 30 

per cent alcohol. 3 parts 

Picric acid, saturated solution in 30 per cent 

alcohol. 1 part 

It would be worth while to try other combinations. 

IODINE 

Iodine is well known as an antiseptic. It is also a good fixing 
agent for unicellular, colonial, and filamentous forms. It penetrates 
rapidly. 

To a saturated solution of potassium iodide in distilled water, 
add iodine to saturation. Filter and dilute with distilled water 
until the solution has a rich brown color. For fixing, dilute still 
further to a light-brown color. The solution fixes in 10 to 24 hours, 
but material may be left in it for several days. Wash thoroughly 
in tap water which has stood long enough to give off all excess of air. 
If the staining of the starch does not disappear, a J per cent solution 
of tannic acid in water will remove any excess color. 

FORMALIN 

Formalin is an excellent preservative. It has been mentioned 
already as an ingredient in several formulas. Commercial formalin has 
a strength of 40 per cent. Throughout this book, a 2,4, or 6 per cent 
formalin is understood to mean 2, 4, or 6 c.c. of commercial for¬ 
malin to 98, 96, or 94 c.c. of water, alcohol, or any other ingredient. 
Commercial formalin is sure to contain some formic acid. For most 
purposes, it is neither necessary nor desirable to remove the acid. 
For studying the origin of vacuoles, it is necessary to have neutral 
formalin, which can be secured from commercial formalin by dis¬ 
tillation. Place some sodium bicarbonate in a flask of formalin and 
distil by heating over a Bunsen flame. It is not worth while to distil 
more than is needed for immediate use, since the formic acid soon 
reappears. 

For filamentous algae and fungi a 3 to 6 per cent solution of the 
ordinary commercial formalin in water is very good. Material is 




32 


METHODS IN PLANT HISTOLOGY 


left in the solution until needed for use. For marine algae sea-water 
should be used instead of fresh water. Both marine and fresh¬ 
water material should be washed for half an hour in fresh water before 
staining. A 6 per cent solution will fix one-fourth its volume of 
material. With material like filamentous algae or leafy liverworts, 
a 10 per cent solution will fix all one can put into the bottle without 
crowding. 

For class use, material should be washed in water for several 
minutes, because the fumes are irritating to the eyes and mucous 
membranes. 

For a study of mitochondria or the origin of vacuoles the following 
combination is recommended: 


Bensley’s Formula. — 

1. Formalin (neutral). 10.0 c.c. 

2. Bichromate of potash. 2.5 g. 

3. Corrosive sublimate. 5.0 g. 

4. Water. 90.0 c.c. 


Make the solution 2, 3, 4, and then add the neutral formalin. 
Fix about 24 hours. Wash in water, but use the iodine—necessary 
on account of the corrosive sublimate—just before staining sections 
on the slide. 

GENERAL HINTS ON FIXING 

It is very desirable that the fixing agent penetrate quickly to all 
parts of the object. For this reason material should be in small 
pieces. The best fixing agents do their best work near the surface 
of the piece. Of course, filamentous algae and fungi, and delicate 
objects like fern prothallia and root-tips, are simply thrown into the 
fixing agent. Alcohol, formalin alcPhol, or formalin alone, may 
penetrate J-inch cubes; but the chromic-acid series, which gives 
the best results in cytological work, penetrates so poorly that cells 
more than X V inch from the surface are not likely to be well fixed. 
Most objects should be trimmed with a razor so that no part shall 
be more than T V inch from the surface. Even then, it must be remem¬ 
bered that a waxy or cutinized or suberized surface presents an almost 
impassable barrier to the chromic series. 

Some objects, although small, cause trouble in various ways. 
Many buds are hairy and will not sink; if such things are dipped 






REAGENTS 


33 


quickly in strong alcohol, they will usually sink. If rather large 
air bubbles prevent the material from sinking, as in case of peri- 
chaetical leaves of some mosses and involucral leaves of liverworts, a 
little dissection or a careful snip with the scissors will obviate the 
difficulty. If an air-pump is available some bubbles are easily 
removed, but air bubbles in cells may resist even the air-pump. 
Heating followed by rapid cooling is recommended by Pfeiffer and 
Wellheim for removing air, but, for cytological work, the remedy is 
worse than the bubbles. 

It is often asked whether fixing agents really preserve the actual 
structure of cell contents. It must be admitted that some things— 
notably the liquid albuminoids—are much modified in appearance, 
but the most competent observers are now inclined to believe that 
such delicate objects as chromosomes, centrosomes, the achromatic 
figure, and even the structure of protoplasm, can be studied with 
confidence from material which has been fixed, imbedded, and 
stained. Extensive investigations upon various objects in the living 
condition have strengthened this confidence. 

It is certain that we have not yet found the ideal fixing agent 
for cell contents. Such an agent must not be a solvent of any of the 
cell contents, must penetrate rapidly, must preserve structures 
perfectly, and must harden so thoroughly that every detail shall 
remain unchanged during the subsequent processes of dehydrating, 
clearing, imbedding, sectioning, and staining. 

DEHYDRATING AGENTS 

Objects which are to be imbedded in paraffin or celloidin, and 
also all other objects which are to be mounted in balsam or Venetian 
turpentine must be dehydrated, i.e., they must be freed from water. 
The slightest trace of water is ruinous. Alcohol is used almost 
exclusively for dehydrating. The process must be gradual. If 
material has been fixed in an aqueous solution, it must pass through 
a series of alcohols of increasing strength, beginning with about 
3 per cent alcohol. Ten years ago, most botanists were beginning with 
35 per cent alcohol; in the second edition of this book (1905) we 
recommended 15, 35, 50, 70, 85, 95, and 100 per cent as a safe series, 
since it causes no obvious plasmolysis of the cell contents. As in¬ 
vestigations have become more and more critical, especially investi¬ 
gations upon the structure of chromatin, it has been found that even 


34 


METHODS IN PLANT HISTOLOGY 


15 per cent alcohol is too strong for a beginning. It is maintained 
that, in addition to the damage done by transferring from water to 
so strong an alcohol, the final dehydration is not so perfect as it is 
when the series begins with a weaker alcohol. Yamanouchi, whose 
work upon delicate algae has been particularly successful, uses the 
following series: 2J, 5, 7§, 10, 15, 20, 30, 40, 50, 70, 85, 95, and 
100 per cent. After such gradual early stages, there seems to be no 
objection to the less gradual stages which follow. Of course, there 
is no particular virtue in the fractions: it is convenient to make a 
10 per cent alcohol, then dilute it one-half for the 5 per cent, and dilute 
the 5 per cent one-half for the 2| per cent. The 7| per cent is made 
with sufficient accuracy by adding a little water to the 10 per cent 
alcohol. For each of the grades up to 50 per cent, one hour is enough 
for objects like fern p^othallia; but an hour and a half should be 
given to each of the remaining grades. Objects like onion root-tips 
should have 2 hours in each of the grades up to 50 per cent, and 3 hours 
in each of the others. Quarter-inch cubes should have 3 or 4 
hours in each grade up to 50 per cent, and 4 or 5 hours in the others. 
The times for larger objects will be longer and will depend upon the 
size and density. In all cases, the absolute alcohol should be changed 
2 or 3 times. The grades below 100 per cent can be used repeatedly. 
The absolute alcohol should not be used again for this purpose, but 
may be put back into the 95 per cent bottle. It is always well to 
filter the alcohols when pouring back into the bottle. Otherwise, 
there would soon be an accumulation of starch grains, pollen grains, 
spores, and various other things. Waste alcohol as strong as 85 or 
95 per cent will be useful for rinsing one’s hands when dealing with 
Venetian turpentine. If it is necessary to be very economical, the 
stronger alcohols may be filtered into a single large bottle and the 
strength of the mixture can then be determined by using an alcohol¬ 
ometer. Knowing the strength of the mixture, one can easily make 
any weaker grade. 

Be very sure that the bottles or Stenders for absolute alcohol 
are perfectly dry; also, keep the bottles well corked and keep 
the lids on the Stenders. The importance of excluding moisture 
cannot be exaggerated. Tightly fitting corks and closely fitting 
covers are better than absorbents; prevention is better than 
cure. 

The lower grades are made up from 95 per cent alcohol. 


REAGENTS 


35 


Formulas for Alcohols.—The following formulas will enable 
anyone to make the other grades of alcohol from 95 per cent alcohol 
and water. 


95 

95 

95 

95 

95 

95 

95 

95 

95 

10 

15 

20 

30 

40 

50 

60 

70 

85 

85 

80 

75 

65 

55 

45 

35 

25 

10 


The foregoing are the formulas for various alcohols from 10 to 
85 per cent. The first column shows the formula for making 10 per 
cent alcohol. The percentage of alcohol secured in each case is 
indicated by the middle number in each column. In the first formula, 
subtract 10from 95; the result, 85, is the number of cubic 
centimeters of water which must be added to 10 c.c. 
of 95 per cent alcohol in order to obtain 10 per cent 
alcohol. The mixture contains 95 c.c. of 10 per cent 
alcohol. If more or less than 95 c.c. of the mixture is 
needed, take proportional parts of 10 and 85. This 
simple method is a time-saver, but if the bottles or 
Stender dishes are to be filled frequently, it will be a 
still further saving of time to use a long label (Fig. 15) 
and, after pouring in the 95 per cent alcohol, draw a 
line showing how high it reaches; and then, after pour¬ 
ing in the water, draw another line. The next time 
it is necessary to fill the bottles merely pour in 95 per 
cent alcohol until it reaches the first line, and then 
pour in water until it reaches the second line. It is 
not necessary to use distilled water if pure drinking-water is available. 

Synthol is used like alcohol, and many believe it to be a good 
substitute. 

Acetone has also been used with more or less success for all 
grades except absolute alcohol. 

Some investigators use more or less complicated diffusion appa¬ 
ratus and make the dehydration process extremely gradual. Judging 
from the finished preparation, we find no advantage in the method. 
In the diffusion process, the solution is constantly changing. This 
may not be an advantage. 

Some very minute objects, like bacteria and the smaller Cyano- 
phyceae, may be dehydrated by heating them until all water is 


70 

per cent 


Fig. 15.— 
Label for 
staining-dish. 







36 


METHODS IN PLANT HISTOLOGY 


drawn off, but, of course, this shows merely the form, with little or 
nothing of the internal structure. 

CLEARING AGENTS 

Clearing agents are so named because they render objects trans¬ 
parent. When clearing agents are used to precede infiltration with 
paraffin, the clearing is merely incidental, the real purpose being to 
replace the dehydrating agent with a solvent of paraffin. The 
clearing is useful, even in this case, because it indicates when the 
replacing has become complete. 

When the clearing agent is used to precede infiltration with 
paraffin, the material should always be most thoroughly dehydrated 
with absolute alcohol before beginning with the clearing agent. 
When the clearing agent is used to clear sections or small objects just 
before mounting in balsam, absolutely perfect dehydration is not 
necessary with all clearing agents. Bergamot oil, carbolic acid, and 
Eycleshymer’s clearing fluid (equal parts of bergamot oil, carbolic 
acid, and cedar oil) will clear readily from 95 per cent alcohol. Sec¬ 
tions to be cleared in xylol or clove oil should be dehydrated in 
“absolute” alcohol. If the absolute alcohol is below 99 per cent, 
xylol will not clear perfectly; but clove oil clears readily from 99 per 
cent and even from 98 or 97 per cent. If the absolute alcohol is 
not well above 99 per cent, it is a good practice to go from the alcohol 
to clove oil; and then, from clove oil to xylol. 

Xylol.—In our opinion, xylol is the best clearing agent to pre¬ 
cede infiltration with paraffin. After the material has been dehy¬ 
drated, it should be brought gradually into xylol. Thirty years 
ago it was customary to bring material directly from absolute alcohol 
into xylol; twenty years ago, two or three mixtures of absolute 
alcohol and xylol were used before reaching the pure xylol; at 
present, those who are doing the most critical work are making this 
process still more gradual. As cytologists have been studying more 
and more minute structures, the methods have become more and 
more critical. As in the case of the alcohol series, the xylol series 
has its grades closer together at the beginning than at the end. 
The following series seems to be sufficiently gradual: T V, J, \ f f, 
pure xylol. It is hardly necessary to use a graduate in making up the 
series. For the J, use equal parts of xylol and absolute alcohol; for 
the i, use equal parts of the \ and absolute alcohol; for the J, use 


REAGENTS 37 

equal parts of the | and absolute, and for the T V, equal parts of the £ 
and absolute. The J can be guessed at with sufficient accuracy. 

Some prefer a closer series of xylols, using 2J, 5, 7|, 10, 15, 20, 
30, 40, 50, 75, and 100 per cent. It is claimed that infiltration with 
paraffin is more thorough after this series. About one hour in each 
grade is enough for filamentous algae and fungi, fern prothallia, and 
similar objects. An hour and a half is enough for onion root-tips. 
For quarter-inch cubes, try 3 hours. For larger pieces the times 
should be longer. In all cases, the pure xylol should be changed 
two or three times. While the pure xylol must not be used again for 
this purpose, it is still good for dissolving paraffin ribbons when 
staining on the slide. 

Xylol is the best agent for clearing sections just before mounting 
in balsam. Preparations cleared in xylol harden more rapidly, 
and this is such a decided advantage that even when sections have 
been cleared in cedar oil or clove oil it is worth while to give them a 
minute or two in xylol before mounting. Besides, clove oil is a solvent 
of many of the most frequently used stains and, consequently, 
preparations in such stains would fade, if transferred directly from 
clove oil to balsam. 

Xylol evaporates so rapidly that one must take care not to let 
sections become dry before applying the balsam. Thin sections 
perfectly dehydrated will clear in a few seconds; but, even with very 
thin sections, it is better to let the xylol act for at least a minute. 
Sections 20 /z in thickness should remain in the xylol 2 to 5 minutes 
before mounting in balsam. If there is much moisture in the air, 
or if the absolute alcohol is not above suspicion, clear sections in 
clove oil before transferring to xylol. 

Chloroform. — Some botanists use chloroform to precede the 
infiltration with paraffin. In the later stages of infiltration it is more 
easily removed than xylol. It seems to possess no other advantages, 
and for clearing sections just before mounting in balsam it is inferior 
to xylol or clove oil. Its value in hardening celloidin and as a fixing 
agent entitles it to a place in the histological laboratory. 

Cedar Oil.—It is not always easy to get good cedar oil. If the 
stuff offered for sale looks like turpentine and smells like it, it is 
worthless for histological purposes. Good cedar oil has a slightly 
amber tint, the color resembling a weak clove oil. It should have 
the pleasant odor of cedar wood. The very expensive cedar oil 


38 


METHODS IN PLANT HISTOLOGY 


used with immersion lenses is not needed for clearing or for preced¬ 
ing infiltration with paraffin. It is claimed that material cleared in 
cedar oil does not become so brittle as that cleared in xylol or chloro¬ 
form. 

Dr. E. J. Krause has used cedar oil extensively in clearing large 
objects—strawberries and gooseberries either whole or cut in two, 
sections of apple 2 to 4 mm. thick, and similar objects. This method 
is proving valuable in vascular anatomy, some material showing the 
course of bundles very clearly in pieces so large as centimeter cubes. 

Xylol can be used in the same way, but is so volatile that speci¬ 
mens often dry up. Dr. Land suggests equal parts of xylol and 
carbon disulphide for clearing large objects which are to be examined 
without sectioning. 

Clove Oil.—This is an excellent agent for clearing sections and 
small objects just before mounting in balsam. It clears more readily 
than xylol. When the absolute alcohol has deteriorated so that xylol 
no longer clears the sections, clove oil may still clear with ease. 
While clove oil will clear from 95 per cent alcohol, it is better to use 
absolute. Since preparations cleared in clove oil harden slowly, it is 
a good plan to treat them with xylol before mounting in balsam. 
Gentian-violet is somewhat soluble in clove oil, and this fact makes 
it possible to secure a beautiful differentiation, because the stain 
is extracted from some elements more rapidly than from others. 
The stain may be extracted completely from the chromosomes 
during the metaphase and still remain bright in the achromatic 
structures. After the desired differentiation has been attained, the 
preparation should be placed in xylol to remove the clove oil, since 
the continued action of the clove oil would cause the preparation to 
fade. Do not use a Stender dish for clove oil, but keep it in a 50 c.c. 
bottle. Put on a few drops, and immediately drain them off in such 
a way as to remove the alcohol as completely as possible. Then 
flood the slide and pour the clove oil back into the bottle, repeating 
the process until the proper differentiation has been reached. Re¬ 
place the clove oil with xylol and mount in balsam. With stains 
not soluble in clove oil, the xylol is not necessary, except to facilitate 
the hardening of the preparation. 

Clove oil may be used in removing the celloidin matrix from 
celloidin sections. It is useless as an agent to precede infiltration with 
paraffin. 


REAGENTS 


39 


Eycleshymer’s Clearing Fluid.—This is a mixture of equal parts 
of bergamot oil, cedar oil, and carbolic acid. It clears readily from 
95 per cent alcohol, and consequently is useful in clearing celloidin 
sections when it is desirable to preserve the celloidin matrix. In 
sections stained with haematoxylin, or haematoxylin and eosin, the 
stain may be removed completely from the matrix by the use of 
acid alcohol, and the matrix may be preserved by clearing from 95 
per cent alcohol. 

It is not intended that the mixture should be used to precede 
infiltration with paraffin. 

Other Clearing Agents.—Bergamot oil, carbolic acid, turpentine, 
benzine, gasoline, and other reagents have been tried for clearing, 
but none seem to be worth more than a warning mention. 

MISCELLANEOUS REAGENTS 

Canada Balsam is used almost exclusively for mounting Very 
thick balsam is disagreeable to handle and makes unsatisfactory 
mounts. Very thin balsam, in drying out, allows bubbles to run 
under the cover. Xylol is cheaper than balsam, and consequently 
the balsam on the market is likely to be too thin for immediate use. 
The stopper may be left out until the balsam acquires the proper 
consistency. Balsam must not be acid. If there is the slightest 
acid reaction, most stains will fade. 

Paraffin should be of at least two grades, a soft paraffin melting 
at 40° to 45° C., and a hard paraffin melting at 52° to 54°C. Griibler’s 
paraffin and most imported paraffins melt at the temperature indicated 
on the wrappers. The melting-point indicated on the wrappers 
of paraffins sold by some American dealers does not enable one to 
make even a guess as to the real melting-point. Paraffin marked 
70° C. may melt at 60° C., and other grades are likely to melt before 
the temperature indicated on the labels is reached. The fact that 
the price rises with the melting-point may explain the discrepancy. 
Test every grade with a thermometer. If it is desired to get a 
paraffin melting at 52° C. and your sample melts at 50° C., add a 
little paraffin with a melting-point above 52° C.; if the sample melts 
at 55° C., add a little with a melting-point below 52° C. 

Paraffin may be used repeatedly. Keeping it in the liquid con¬ 
dition in the bath month after month is an advantage, since it 
becomes more and more tenacious and homogeneous. 


40 


METHODS IN PLANT HISTOLOGY 


Glycerin, glycerin jelly, Venetian turpentine, and gold size are 
described in the chapter on “The Glycerin Method” (chap. vii). 
Celloidin is described in the chapter on “The Celloidin Method” 
(chap, x), and cellulose acetate in the chapter on “The Cellulose 
Acetate Method” (chap. xi). The reagents already described are 
noted further in connection with specific applications. Reagents 
used in making microchemical tests are described in the chapter on 
“Temporary Mounts and Microchemical Tests” (chap. v). 

A list of reagents with suggestions in regard to quantities and 
prices will be found in chapter xxix. 


CHAPTER III 


STAINS AND STAINING 

Since the third edition of this book appeared in 1915 no new 
stains of the first rank have come into favor, but much greater 
precision has been attained in the use of some which were already 
popular. For cytological work Haidenhain’s iron-haematoxylin 
holds a firm place at the head of the list, with Flemming’s triple 
stain an easy second. For anatomical work, safranin still holds 
first place for the lignified elements of the vascular system, but the 
claim of Delafield’s haematoxylin to first place for cellulose tissues 
is no longer undisputed, for anilin blue is giving excellent results and 
light green seems to give more accurate views of the phloem than we 
were securing with any of the other stains. The fact that excellent 
preparations can be made, almost without trial, by using combina¬ 
tions already perfected doubtless deters investigators from experi¬ 
menting with other stains. There is still abundant room for experi¬ 
menting with various stains, and especially in the use of mordants and 
in the effect of the same stain or combination after various fixing 
agents. It is to be regretted that botanists who need microtechnic 
have so little knowledge of chemistry, and that chemists have no 
interest in developing methods of staining. During the past few 
years, American stains have been developed until many equal and 
some even surpass the famous Griibler products; and, besides, the 
American stains are becoming standardized. 

Stains may be classified in various ways: e.g., there are three 
great groups of stains—the carmines, the haematoxylins, and the 
anilins. Stains may be classified as basic and acid, or they may be 
regarded as general and specific. A general stain affects all the 
elements, while a specific stain affects only certain elements, or 
stains some elements more deeply than others. Stains which 
show a vigorous affinity for the nucleus have been called “nuclear 
stains,” and those which affect the cytoplasm more than the nucleus 
have been termed “plasma stains.” Of course, such stains are 
specific. 


41 


42 


METHODS IN PLANT HISTOLOGY 


We shall consider some of the more important haematoxylins, 
carmines, and anilins, reserving general directions and theoretical 
questions for another chapter. The formulas are largely empirical. 
Some of those given here are taken from The MicrotomisVs Vade- 
Mecum (Lee), which is easily the most complete compendium of 
stains and other reagents concerned in microtechnic. It is to be 
regretted that botanists have no book of this character, but it must 
be confessed that we have not the material for such an extensive 
work. Other formulas are from Botanical Microtechnique (Zimmer- 
mann) and from Stirling’s Histology , and still others are from current 
literature and from our own laboratory. The directions for using a 
stain apply to stains made up according to the formulas which are 
given here, and may need modification if other formulas are employed. 
It is hoped, however, that the directions will give the student sufficient 
insight into the rationale of staining to enable him to make any 
necessary modifications. Since American stains have come into 
general use, the need for rationale is even greater, especially if the 
American stains are made up according to standard formulas, which 
are based largely upon the Griibler products. In general, it would 
seem that the American stains are purer and that they act more 
rapidly. 

The current practice in staining paraffin sections on the slide 
differs from the practice in staining freehand sections or small objects 
which are to be mounted whole. In case of paraffin sections, the 
cell contents are usually as important and often more important 
than the cell walls; consequently, extreme care must be given to every 
detail. With freehand sections the cell contents often drop out, 
but even when they remain, the cell walls are usually the important 
features; and so the process is considerably shortened. 

For staining freehand sections, it is customary to use solid watch 
glasses, unless the sections are very large. The details of the method 
are given in chapter vi, on “Freehand Sections.” 

For staining sections on the slide, nothing is better than the 
ordinary Stender dish. The arrangement of Stender dishes shown 
in Figure 16 is very convenient. The advantage is obvious. With 
two dishes each of xylol, xylol-alchol, and absolute alcohol, one 
set can be used in passing down to the stain, and the other, which 
is thus kept free from any paraffin in solution, can be used in passing 
back to the balsam. Even for paraffin sections, some use only 


STAINS AND STAINING 


43 


three alcohols, 50, 95 and 100 per cent, and the first two may be 
simply poured over the slide; in this case, only one Stender dish— 
for the 100 per cent alcohol—is necessary in the alcohol series, the 
other two alcohols being kept in bottles. This short method gained 
great popularity because it was used in Strasburger’s laboratory at 
Bonn. It was the influence of this school and its great master 
which led to the adoption of the short schedule in the second edition 



of this book. A few years’ trial showed the weakness of the method, 
and we returned to the longer schedule. The crudeness of the 
short schedule is doubtless responsible for the tenacity with which 
the Bonn school has clung to the theory of linin and chromomeres. 
The young investigator should be warned that during the last twenty 
years of his life, Strasburger, who had been a leader in technic, cut 
very few sections and did practically no staining, but used prepara¬ 
tions made by assistants. 

Let us now consider a few of the most important stains. 

THE HAEMATOXYLINS 

The most important haematoxylins are Haidenhain’s iron-alum 
haematoxylin, Delafield’s haematoxylin, Mayer’s haem-alum, and 
Boehmer’s haematoxylip. 

All the haematoxylins mentioned contain alum, and, according 
to Mayer, who has written the most important work on haematoxylin 
stains, 1 “the active agent in them is a compound of haematin with 
alumina. This salt is precipitated in the tissues, chiefly in the nuclei, 
by organic and inorganic salts there present (e.g., by the phosphates), 
and perhaps also by other organic bodies belonging to the tissues.” 
These salts are fixed in the tissues by the killing and fixing agent, 
and when the stain is applied a chemical combination results. 
Haematoxylins stain well after any of the fixing agents described in 

1 “Ueber das Farben mit Hamatoxylin,” Mittheilungen aus der Zoologischen Station zu Neapel, 
io:170-186, 1891, and “Ueber Hamatoxylin, Carmin und verwandte Materien,” Zeitschrift fiir 
mssenschaftliche Mikroscopie, 16:196-220, 1899. 






44 


METHODS IN PLANT HISTOLOGY 


the preceding chapter, but they are most effective when used after 
members of the chromic-acid series. 

Haidenhain’s Iron-Alum Haematoxylin. — This stain, introduced 
by Haidenhain in 1892 immediately gained great popularity and now, 
after more than 30 years’ constant use, still maintains first place in 
cytological investigations. Two solutions are used, and they are 
never mixed: 

A. 2 to 4 per cent aqueous solution of ammonia sulphate of iron. 

B. | per cent aqueous solution of haematoxylin. 

In making solution A, use the violet ferric crystals, hot the ferrous. 

The first solution acts as a mordant, i.e., it does not stain, but 
prepares the tissue for the action of the second solution. 

Solution A is at its best as soon as the crystals are completely 
dissolved and it remains in practically perfect condition for about 
two months, after which it gradually deteriorates. 

The haematoxylin crystals for solution B should be dissolved in 
water.. This will require about 10 days. The solution should then 
be allowed to “ripen” for 4 weeks before it is ready for use. Unfor¬ 
tunately, it remains at its best for only a short time, not more than 
5 or 6 weeks. This is because the “ripening,” which is an oxidation 
process, continues, and the solution becomes too ripe. During the 
ripening, a cotton plug should be used instead of a cork, to facilitate 
the oxidation; but as soon as the stain is ripe, a cork—preferably 
a perfectly fitting glass stopper—will prolong the maximum efficiency. 
Some prefer to dissolve the haematoxylin crystals in alcohol—-about 
10 g. in 100 c.c. of absolute alcohol. This solution should stand until 
it has a deep wine-red color. This will require 4 or 5 months, and a 
year is not too long. From this stock solution, make up small quanti¬ 
ties as needed. About 4 or 5 c.c. of this stock solution in 100 c.c. of 
water gives a practically aqueous solution, and it is already ripe. 

The American haematoxylin ripens faster and may reach its 
maximum efficiency in 2 or 3 weeks. One might naturally expect 
that it would retain its efficiency for a correspondingly shorter time; 
but we have found that, even for the structure of chromatin, the stain 
seems to stay at its best for at least two months. 

The general method is as follows: treat with A, stain in B, and 
then return to A to reduce and differentiate the stain. Never 
transfer directly from A to B, or from B to A; always wash in water 
before passing from one of the solutions to the other. 


STAINS AND STAINING 


45 


While all follow the general method just indicated, no two investi¬ 
gators would prepare exactly the same schedule, even for staining the 
same object, e.g., root-tips; neither investigator would use the same 
schedule for a root-tip and an embryo sac; an alga might require 
different treatment, and all the preceding variations might fail 
miserably with the pollen tubes of cycads. This stain is so important 
that every worker must learn it, and the only way to learn it is to 
become acquainted with the general outline of the process and then 
adapt every step to the case in hand. 

For the sake of illustration, I asked two prominent cytologists, 
Dr. S. Yamanouchi and Dr. L. W. Sharp, both of whom have been 
notably successful in staining mitotic figures, to write schedules 
indicating their methods of using this stain. While both protested 
that the practice could not be written down, they kindly prepared 
the following schedules, not for the instruction of their colleagues, but 
to introduce the method to beginners. Both schedules are for par¬ 
affin sections. Throughout the first schedule, I have interpolated 
comments and suggestions. 

Yamanouchi’s Schedule.— 

1. Xylol, 5 minutes, to dissolve the paraffin. 

Do not heat the slides to melt the paraffin. However, a gentle 
warming which does not approach the melting-point of the paraffin does 
no damage and makes the paraffin dissolve more readily. The xylol 
soon has considerable paraffin in solution, but 100 c.c. of xylol should 
remove the paraffin from at least 100 slides with ribbons 25 mm.long and 
10 ix thick. If the ribbons are only 5 \x thick, 200 slides can be treated. 

2. Xylol and absolute alcohol, equal parts, 5 minutes. 

3. Absolute alcohol, 5 to 7 minutes. 

4. 95, 85, 70, 50, 35 per cent alcohol, 5 minutes each. 

If material has been fixed in a reagent containing osmic acid, it 
should be bleached. For this purpose, 10 to 15 c.c. of hydrogen peroxide 
may be added to 100 c.c. of the 50 per cent alcohol. 

5. Water, 10 to 20 minutes. 

If any alcohol is left in the sections, the staining will not be brilliant. 
Change the water several times. 

6. Iron-alum. 

Use the 4 per cent solution. For many objects, like the archegonia 
of gymnosperms and the embryo sacs of angiosperms, 1 hour is usually 
enough. For chromosomes in root-tips and anthers, 2 hours may be 
long enough; but for algae, 2 hours is generally a minimum. 


46 


METHODS IN PLANT HISTOLOGY 


7. Wash in water, 5 minutes. 

The water should be changed several times. If the washing is not 
thorough, the differentiation will not be sharp. 

8. Haematoxylin. 

Many objects, like the archegonia of gymnosperms and the embryo 
sacs of angiosperms, will stain sufficiently in 5 or 6 hours; most algae 
require at least 20 hours. 

9. Wash in water, 5 minutes, changing as often as the water shows any color. 

10. Iron-alum, 2 per cent solution. 

No time can be indicated here. The preparation must be watched 
under the microscope. After some experience, on'e can form some 
judgment from the color tone, as the slide stands in the Stender dish 
of iron-alum, but the finishing must always be done under the microscope. 
If the stain is coming out rather slowly, as it should, one can handle 
6 to 10 slides at one time. Put the slides on a 5X7 glass plate and put 
the plate on the stage of the microscope. The iron-alum can be added 
or removed with a pipette. As slide after slide reaches the proper 
differentiation, it is placed in water. 

11. Water, 30 minutes. 

The water should be changed several times. If this washing is 
not thorough, the preparation will fade, on account of the continued 
action of the iron-alum. If an aqueous counter-stain is used, apply it 
at this point. 

12. 35, 50, 70, 85, 95, 100 per cent alcohol, 5 minutes in each. 

If an alcoholic counter-stain is used, apply it near the alcohol of 
the same strength as the stain. 

13. Absolute alcohol and xylol, equal parts, 5 minutes. 

14. Xylol, 2 to 5 minutes. 

15. Balsam. 

Sharp’s Schedule.— 

1. Remove the paraffin with xylol. 

2. Rinse in absolute alcohol. 

3. 95 per cent alcohol. 

4. 50 per cent alcohol. 

5. Water. 

6. If osmic acid has been used in fixing, place the slides in 10 per cent 
solution of peroxide of hydrogen in water until bleached. 

7. Water. 

8. Iron-alum, 2§ per cent, 2 to 3 hours. 

9. Wash well. 

10. ^ per cent haematoxylin, 24 hours. 

11. Wash in water. 


STAINS AND STAINING 


47 


12. Extract the stain in 1 per cent iron-alum, watching the process under the 
microscope. 

13. Wash several hours in-water. 

14. Alcohol series: 10, 30, 50, 70, 80, 95, 100 per cent. 

If a counter-stain is desired, introduce it in one of the alcohols of 
this series. 

15. Absolute alcohol and xylol, equal parts. 

16. Xylol. 

17. Mount in balsam. 

While these two schedules would enable the student to apply 
the method in case of objects to be mounted whole, like filamentous 
algae, fern prothallia, etc., a complete schedule is given in chapter 
viii on “The Venetian Turpentine Method.” 

The times given above must not be accepted as final. Many 
prefer to wash in water for several hours after the first immersion in 
iron-alum. Some think that 4 hours is enough for the entire process. 
Many put the slide into iron-alum in the morning and finish the pro¬ 
cess in the afternoon. These short schedules are not likely to prove 
satisfactory with mitotic figures. A plan which has proved con¬ 
venient and very successful is to put the slide into the iron-alum 
in the morning, wash in water for an hour at some convenient 
time in the afternoon, leave it in the § per cent haematoxylin over 
night and finish the preparation the next morning. It is a long 
process, requiring care, patience, and judgment, but it is worth the 
effort. 

Chromosomes, centrosomes, and pyrenoids take a brilliant black; 
or, if the second treatment with iron-alum be more prolonged, 
a blue black or purple. Achromatic structures stain purple, but 
the stain can be extracted while it is still bright in the chromosomes. 
Lignified, suberized, and cutinized structures stain lightly or not at 
all. Cellulose does not stain so deeply as with Delafield’s haema¬ 
toxylin. Archesporial cells and early stages in sporogenous tissue 
stain gray. Many details which are not so brilliantly colored often 
show good definition. 

If a counter-stain is desired, anything which gives a serviceable 
contrast may be used. In any case, the haematoxylin stain must be 
complete and the washing thorough before the second stain is applied. 
An aqueous stain should be applied just after the final washing in 
water; an alcoholic stain should be applied during the process of 


48 


METHODS IN PLANT HISTOLOGY 


passing the slides through the alcohols, staining in a solution of saf- 
ranin in 50 per cent alcohol from the alcohol of a concentration 
nearest that of the stain; and staining after the final absolute alcohol, 
if the stain is dissolved in clove oil. 

A stain of 3 or 4 minutes in safranin adds an excellent differentia¬ 
tion in case of many algae and does not obscure nuclear details. 
The exine of pollen grains may take a brilliant red with safranin 
in 5 or 10 minutes, contrasting sharply with the mouse gray of the 
intine. Orange G, in clove oil, often gives a pleasing contrast. 

Delafield’s Haematoxylin. — “To 100 c.c. of a saturated solution 
of ammonia alum add, drop by drop, a solution of 1 g. of haema¬ 
toxylin dissolved in 6 c.c. of absolute alcohol. Expose to air and 
light for one week. Filter. Add 25 c.c. of glycerin and 25 c.c. of 
methyl alcohol. Allow to stand until the color is sufficiently dark. 
Filter, and keep in a tightly stoppered bottle” (Stirling and Lee). 
The addition of the glycerin and methyl alcohol will precipitate 
some of the ammonia alum in the form of small crystals. The last 
filtering should take place 4 or 5 hours after the addition of the 
glycerin and methyl alcohol. 

The solution should stand for at least two months before it is 
ready for using. This “ripening” is brought about by the oxida¬ 
tion of haematoxylin into haematin, a reaction which may be secured 
in a few minutes by a judicious application of peroxide of hydrogen. 
However, we prefer to let the haematoxylin ripen naturally. There 
is no objection to making this stain in considerable quantity, since 
it does not deteriorate. We have used Delafield’s haematoxylin 
which had been in a cork-stoppered bottle for twenty years, and 
it still gave the rich characteristic stain. 

Transfer to the stain from 50 or 35 per cent alcohol or from water. 
The length of time required is exceedingly variable. Sometimes 
sections will stain deeply in 3 minutes, but it is often necessary to 
stain for 30 minutes or even longer. This stain may be diluted 
with several times its own volume of water; when this is done, the 
time required is correspondingly long, but the staining is frequently 
more precise. The length of time required will be fairly uniform for 
all material taken from the same bottle. This fact indicates that the 
washing process, which follows killing and fixing, is an important 
factor; if the washing has been thorough, the material will stain 
readily; but if the washing has been insufficient, the material may 


STAINS AND STAINING 


49 


stain slowly or not at all. The washing is particularly important 
when the fixing agent contains an acid. Transfer from the stain 
to tap water. Distilled water is neither necessary nor desirable. 
Some writers recommend washing for 24 hours, but this is entirely 
unnecessary; for paraffin sections on the slide, 5 or 10 minutes is 
long enough, and even for rather thick freehand sections 20 or 30 
minutes is sufficient. Use plenty of water and keep changing it as 
often as it becomes in the least discolored. Precipitates are often 
formed when slides are transferred directly to alcohol from this 
stain, and sometimes even after washing in water. A few gentle 
dips in acid alcohol (2 drops of HC1 to 100 c.c. of 70 per cent alcohol) 
will usually remove the precipitates. This extracts the stain more 
rapidly from other parts than from the nuclei, and hence gives a good 
nuclear stain, while at the same time it removes any disfiguring 
precipitates. Some prefer to stain for a very short time and use no 
acid alcohol, but, as a rule, it is better to overstain and then differen¬ 
tiate in this way, because sharper contrasts are obtained. Transfer 
from acid alcohol to 70 per cent alcohol and leave here until a rich 
purple color replaces the red due to the acid. Since small quantities 
of the acid alcohol are carried over into the 70 per cent alcohol, it 
is well to add a drop of ammonia now and then to neutralize the 
effect of the acid. Too much ammonia is to be avoided, for it gives 
a disagreeable bluish color with poor differentiation, probably on 
account of the precipitation of alumina. The preparation is now 
dehydrated in 95 per cent and then in absolute alcohol, cleared in 
xylol or clove oil, and mounted in balsam. 

The following is a general schedule for staining paraffin sections 
on the slide in Delafield’s haematoxylin: 

1. Stain (from water or from 35 or 50 per cent alcohol) 10 minutes 


2. Rinse in water. 10 minutes 

3. 35 and 50 per cent alcohol. 3 minutes each 

4. Acid alcohol. 5 seconds 

5. 70 per cent alcohol. 3 minutes 

6. 85 per cent alcohol. 3 minutes 

7. 95 per cent and 100 per cent alcohol.3 minutes each 

8. Xylol and 100 per cent alcohol, equal parts. 3 minutes 

9. Xylol. 3 minutes 

10. Mount in balsam. 










50 


METHODS IN PLANT HISTOLOGY 


If, after rinsing in water, the stain is evidently too weak, put 
the slide or section back into the stain until it appears overstained. 
Place the slide in acid alcohol. If an acid alcohol with 2 drops of 
HC1 to 100 c.c. of 70 per cent alcohol reduces the stain too much in 4 
or 5 seconds, use less acid or stain longer. Transfer to 70 per cent 
alcohol without any acid. As soon as the color changes from red to 
purple, examine under the microscope. If it is still overstained, 
return to the acid alcohol; if the stain is too weak, return to the 
haematoxylin and try it again. After the haematoxylin is just right, 
apply a contrast stain, if you wish to double stain. Before trans¬ 
ferring to the xylol wipe the alcohol from the back of the slide, or at 
least rest the corner of the slide upon blotting-paper for two or three 
seconds, in order that you may not carry over so much alcohol into 
the xylol. Add a drop of balsam and a cover. Since the xylol is very 
volatile, this last step must be taken quickly. If blackish spots 
appear they are usually caused by the drying of sections before the 
balsam and cover are added; if there are whitish spots or an emulsion¬ 
like appearance, the clearing is not thorough; this may be caused by 
poor xylol (or other clearing agent); by absolute alcohol which is 
considerably weaker than its name implies (the absolute alcohol 
must test at least as high as 99 per cent, and ought to test as high 
as 99.5 per cent, if xylol is to be used for clearing); or by passing 
too quickly through the absolute alcohol and xylol, or even by 
moisture on the cover-glass. The last danger is easily avoided by 
passing the cover quickly through a Bunsen or alcohol flame before 
laying it on the balsam. 

Delafield’s haematoxylin is the most generally useful stain in the 
haematoxylin group. It brings out cellulose walls very sharply, 
and consequently is a good stain for embryos and the fundamental 
tissue system in general. With safranin it forms a good combination 
for the vascular system, the safranin giving the lignified elements a 
bright red color, while the haematoxylin stains the cellulose a rich 
purple. It is a good stain for chromatin, and the achromatic struc¬ 
tures show up fairly well, but can be brought out much better by 
special methods. Archesporial cells and sporogenous tissue are 
very well defined if proper care be taken. Lignified and suberized 
walls and also starch and chromatophores stain lightly or not at all. 
Whenever you are in doubt as to the selection of a stain for general 
purposes, we should advise the use of Delafield’s haematoxylin. 


STAINS AND STAINING 


51 


Mayer’s Haem-Alum. — Haematoxylin, 1 g., dissolved with gentle 
heat in 50 c.c. of 95 per cent alcohol and added to a solution of 50 g. 
of alum in a liter of distilled water. Allow the mixture to cool and 
settle; filter; add a crystal of thymol to preserve from mold (Lee). 

It is ready for use as soon as made up. Unless attacked by mold, 
it keeps indefinitely. Transfer to the stain from water. It is seldom 
necessary to stain for more than 10 minutes, and 4 or 5 minutes 
is generally long enough. As a rule, better results are secured by 
diluting the stain (about 1 c.c. to 10 c.c. of distilled water) and 
allowing it to act for 10 hours or over night. 

This is a good stain for the nuclei of filamentous algae and fungi, 
since it has little or no effect upon cell walls or plastids. Wash 
thoroughly in water, transfer to 10 per cent glycerin, and follow the 
Venetian turpentine method, as described in chapter viii. 

Erlich’s Haematoxylin.— 


Distilled water. 50 c.c. 

Absolute alcohol. 50 c.c. 

Glycerin. 50 c.c. 

Glacial acetic acid. 5c.c. 

Haematoxylin. 1 g. 

Alum in excess. 


Keep it in a dark place until the color becomes a deep red. If 
well stoppered, it will keep indefinitely. Transfer to the stain from 
50 per cent or 35 per cent alcohol. Stain 5 to 30 minutes. Since 
there is no danger from precipitates and the solution does not over¬ 
stain, it is not necessary to treat with water or with acid alcohol, but 
the slide may be transferred from the stain to 70 per cent alcohol. 
Eosin, erythrosin, or orange G are good contrast stains. Jeffrey 
uses safranin and Erlich’s haematoxylin for woody tissues. 


Boehmer’s Haematoxylin.— 

^ ( Haematoxylin. 1 g. 

\ Absolute alcohol. 12 c.c. 

/ Alum. 1 g. 

1 Distilled water. 240 c.c. 


The solution A must ripen for two months. When wanted for 
use, add about 10 drops of A to 10 c.c. of B. Stain 10 to 20 minutes. 
Wash in water and proceed as usual. 











52 


METHODS IN PLANT HISTOLOGY 


Cellulose walls take a deep violet. The closing membrane 
(torus) of the bordered pits of conifers will usually stain deeply in 
about 15 minutes. Lignified, suberized, and cutinized structures 
stain slightly or not at all. When they do stain, the color is not 
violet, but a light yellow or brown. 

THE CARMINES 

Botanists have never given the carmines a fair trial, doubtless 
because the stains were not considered worth it; but the splendid 
preparations by Professor Powers of various merfibers of the Volvo- 
caceae prove that we should pay more attention to this group. Only 
a few of the multitudinous formulas will be considered. 

The carmine solutions keep for several years, some of them even 
improving with age, if distilled water has been used in the formulas 
and the stains have been kept in tightly stoppered bottles. If the 
solution becomes turbid, it should be filtered. 

Greenacher’s Borax Carmine.— 


Carmine... 3 g. 

Borax.-. 4 g. 

Distilled water. 100 c.c. 


Dissolve the borax in water and add the carmine, which is 
quickly dissolved with the aid of gentle heat. Add 100 c.c. of 70 
per cent alcohol and filter (Stirling). 

The following is a slightly different method for making this stain 
from the ingredients mentioned above: Dissolve the borax in water, 
add the carmine, and heat gently for 10 minutes; after the solution 
cools, add the alcohol and filter; let the solution stand for 2 or 3 weeks, 
then decant and filter again. 

Stain the material in bulk from 50 per cent alcohol 1 to 3 days, 
then treat with acid alcohol (50 c.c. of 70 per cent alcohol+2 drops 
of hydrochloric acid) until the color becomes a clear red; this may 
require only a few hours, but may take 2 or 3 days. The material 
may then be passed through the rest of the alcohols (6 to 24 hours 
each), cleared, imbedded, and cut. After the sections are fastened 
to the slide, the paraffin should be dissolved off with xylol. The 
balsam and cover may be added immediately, or the xylol may be 
rinsed off with alcohol and a contrast stain may be added. 





STAINS AND STAINING 


53 


Alum Carmine. —A 4 per cent aqueous solution of ammonia alum 
is boiled 20 minutes with 1 per cent of powdered carmine. Filter 
after it cools (Lee). 

Stain from 12 to 24 hours and wash in water. No acid alcohol 
is needed, since the solution does not overstain. 

Carmalum (Alum Lake). —Use 1 gram of the powdered stain to 
100 c.c. of very dilute ammonia water. Filter, if there is any 
precipitate. 

Mayer’s Carmalum.— 


Carminic acid. 1 g. 

Alum. 10 g. 

Distilled water. 200 c.c. 


Dissolve with heat; decant or filter and add a crystal of thymol 
to avoid mold. 


Alum Cochineal. — 

Powdered cochineal. 50 g. 

Alum. 5 g. 

Distilled water. 500 c.c. 


Dissolve the alum in water, add the cochineal, and boil; evapo¬ 
rate down to two-thirds of the original volume, and filter. Add a 
few drops of carbolic acid to prevent mold (Stirling). 

Stain as with alum carmine. It used to be a common practice 
to stain in bulk in alum cochineal and counter-stain on the slide 
with Bismarck brown. 

Iron Aceto-Carmine 1. —For counting chromosomes in pollen 
mother-cells mounted whole, Belling and also Sand used a modified 
aceto-carmine method. The preparations are good for an immediate 
count, but do not last longer than a few days or a week. 

“ Ordinary aceto-carmine is prepared by heating a 45 per cent 
solution of glacial acetic acid to boiling with excess of powdered 
carmine, cooling, and filtering. The young anthers are teased out 
with steel blades or needles in a drop of this until it changes slightly 
toward bluish red. An excess of iron spoils the preparation. 

Iron Aceto-Carmine. — “To a quantity of aceto-carmine a trace 
of solution of ferric hydrate dissolved in 45 per cent acetic acid is 
added until the liquid becomes bluish red, but no visible precipitate 
forms. An equal amount of ordinary aceto-carmine is then added. 








54 


METHODS IN PLANT HISTOLOGY 


The anthers are teased out with nickled instruments.” A cover- 
glass is then added and sealed with vaseline. The preparation lasts 
only a few days, but is much superior to any obtained by the usual 
intra vitam processes. 

THE ANILINS 

Many of the most brilliant and beautiful stains yet discovered 
belong to this group. These stains are very numerous, but not so 
numerous as their names; for different names have been given to 
the same stain, and the same name has been given to different stains. 
Fortunately, the Committee on Standardization of Biological Stains 
is doing a good work in standardizing the nomenclature as well as 
the stains themselves. A valuable list of synonyms, with the pre¬ 
ferred designations, was published in Science, 57:743-746, 1923, 
and other references to the work of the commission are given in the 
Bibliography on page 341. 

General Formula.—Make a 10 per cent solution of anilin oil in 
95 per cent alcohol; when the anilin oil is dissolved, add enough 
water to make the whole mixture about 20 per cent alcohol; add 1 g. 
of cyanin, erythrosin, safranin, gentian-violet, etc., to each 100 c.c. of 
this solution. Solutions containing anilin oil do not keep as well as 
aqueous or alcoholic solutions. 

The anilins keep well in balsam, but not so well in glycerine. 
Xylol is a good clearing agent for all of them; but clearing in clove 
oil improves stains like gentian-violet, which are more or less soluble 
in clove oil. Even in such cases, xylol should follow the clove oil, 
or the preparation will fade. 

While the anilins are not as permanent as the haematoxylins, 
most of them keep fairly well if the staining has been carefully done. 
Preparations fade if exposed long to bright sunlight. Keep the slides 
in the box when not in use, and even when in use, do not leave them 
on the laboratory table, exposed to the sun. We have preparations, 
made more than 25 years ago, in which the safranin and gentian- 
violet are still bright; and others made more than 10 years ago, in 
which Magdala red and aniline blue have not faded. 

Some of the anilins are acid, some basic, and some are neutral. 

The rapidity with which sections must be transferred from one 
fluid to another makes many of them more difficult to manage than 
the haematoxylins or the carmines, but the stains are so valuable 
that even the beginner should spend most of his time with the anilins. 


STAINS AND STAINING 


55 


Many anilins stain quite deeply in 1 to 20 minutes, but if the 
stain washes out during the dehydrating process, stain longer, even 
10 to 24 hours if necessary. Often the brilliancy of the stain can be 
increased by leaving the slide for 5 minutes in a 1 per cent solution of 
permanganate of potassium before staining. The permanganate acts 
as a mordant. 

The following are the more important anilins now in use by botan¬ 
ists. The directions apply to solutions made up according to the 
formulas given with the different stains. 

Safranin. — Two safranins are sold by dealers, one soluble in 
water and the other soluble in alcohol. The alcoholic is somewhat 
soluble in water and the aqueous is somewhat soluble in alcohol, but 
both make better solutions when used with their intended solvents. 

The best aqueous solution is simply a 1 per cent solution in 
distilled water. 

The alcoholic solution is made by dissolving 1 g. of the alcoholic 
safranin in 100 c.c. of 95 per cent or absolute alcohol and, after the 
1 safranin is completely dissolved, adding 50 c.c. of distilled water. 

According to Flemming, dissolve 0.5 g. of alcoholic safranin in 
50 c.c. of absolute alcohol, and after 4 days add 10 c.c. of distilled 
water. 

A method which we have used for many years with good results 
is to make a 1 per cent solution of the aqueous safranin in distilled 
water; then make a 1 per cent solution of the alcoholic safranin in 
95 per cent alcohol; then mix equal volumes of the two solutions. 
This makes a strong solution of safranin in about 50 per cent alcohol. 

The first American safranins were very unsatisfactory but there 
have been great improvements and one company, the National 
Anilin and Chemical Company, has produced a safranin with 90 per 
cent dye content, much stronger than any of the European stains. 
This is the safranin which has been certified by the Committee on 
Standardization of Stains. Other companies are also improving. 
Coleman and Bell's Safranin Y, for bacilli, is good for staining xylem. 

All safranins keep indefinitely, solutions 20 years old staining 
as well, or better, than when fresh. 

An anilin safranin may be made according to the general formula. 

The transfer to the stain depends upon the formula. If the stain 
is aqueous, transfer to the stain from water; if made up according to 
the general anilin oil formula, transfer to the stain from water or, 


56 


METHODS IN PLANT HISTOLOGY 


if coming down from higher alcohols, from 35 per cent alcohol; if 
the mixture of aqueous and alcoholic safranins is used, transfer from 
35 per cent alcohol, if going up in the series, and from 70 per cent 
alcohol, if coming down from stronger alcohols. For freehand 
sections of woody tissues we always use the mixture. If sections are 
cut from living material, leave them in 95 per cent alcohol for half 
an hour and then transfer to the stain. Sections cut from alcoholic 
material may be transferred directly to the stain. If cut from 
formalin-alcohol material, leave the sections in 50 per cent or 70 
per cent alcohol for ten minutes before transferring. If cut from 
formalin material, leave them in water for 10 minutes, then in 35 
per cent alcohol for 10 minutes before staining. 

The time required for staining varies with the tissue, the fixing 
agent, and the quality of the stain. In general, it may be said that 

2 hours is a minimum and 24 hours a maximum. If the staining be 
too prolonged, delicate structures, like starch grains, crystals, and 
various cell constituents, may wash out. The mere fact that the 
whole section does not wash off does not mean that everything is 
fastened to the slide. On the other hand, with a short period, it is 
difficult to get a sharp differentiation. In staining a vascular bundle, 
one should be able to wash the safranin from the cellulose walls and 
still leave a brilliant red in lignified structures. For paraffin sections, 

3 to 6 hours will usually be sufficient. It is a good practice to put the 
slides into the stain in the morning and finish the mounts any time 
in the afternoon. For freehand sections of woody tissues, 24 hours 
is not too long. 

From the stain transfer to 50 per cent alcohol. If the sections 
are deeply stained, and sufficient differentiation is not secured within 
5 or 10 minutes, a drop of hydrochloric acid added to 50 c.c. of the 
alcohol will hasten the extraction of the stain. If staining vascular 
tissue, draw the stain from the cellulose walls, but stop before the 
lignified walls begin to fade. If a contrast stain is to be added, 
like light green, which weakens the safranin; or aniline blue or 
Delafield’s haematoxylin, which need to be followed by an acid; 
the safanin should be strong enough to allow the necessary reduction. 
If staining mitotic figures, draw the stain from the spindle, but stop 
before the chromosomes begin to weaken. When the desired differ¬ 
entiation has been reached, wash out the acid in 50 per cent alcohol, 
if acid has been used. About 5 minutes should be sufficient. 


STAINS AND STAINING 


57 


If safranin is to be used alone, pass through 50, 70, 85, 95, and 
100 per cent alcohol, through the xylol-alcohol, then through xylol 
to balsam. If clove oil is used, omit the xylol-alcohol, but follow 
the clove oil with xylol to hasten the hardening of the preparation. 

If a second stain is to be added, transfer from the 50 per cent 
alcohol to any alcoholic stain. If the second stain is an aqueous 
stain, rinse the slide or sections for a minute in water before applying 
the stain. 

Safranin is the most generally useful of all the red stains, and, 
fortunately, it is quite durable. Lignified, suberized, cutinized, and 
chitinized structures stain red, as do also the chromosomes, nucleoli, 
and centrosomes. 

Acid Fuchsin.—Use a 1 per cent solution in water or in 70 per cent 
alcohol. The solution in alcohol is preferable if sections are to be 
mounted in balsam. This stain often acts with great rapidity, 
2 or 3 minutes being sufficient. The method for using acid fuchsin 
with woody tissues is given in the chapter on “Freehand Sections’’ 
(chap. vi). In staining embryo sacs, pollen grains, and such struc¬ 
tures, longer periods are better. Stain 1 or 2 hours, and then differ¬ 
entiate in a saturated solution of picric acid in 70 per cent alcohol. 
This may require 30 seconds, or even several minutes. Rinse in 
70 per cent alcohol until a bright red replaces the yellowish color 
due to the acid, and then proceed as usual. 

Congo Red.—This is an acid stain resembling acid fuchsin. 
For cytological work use a \ per cent aqueous solution; for anatomical 
work use a saturated solution. It is a good stain to use after mala¬ 
chite green or anilin blue. Transfer to the Congo red from water, 
stain 15 minutes, wash in water, transfer—for wood sections—to 
85 per cent alcohol, and wash until the green or blue color of the 
previous stain begins to show through the red. Then treat quickly 
with absolute alcohol, clear xylol, and mount in balsam. 

Eosin.—This has long been a favorite stain, but for most pur¬ 
poses it has been replaced by similar stains giving better differentia¬ 
tion. The dry stain is made in two forms, one for aqueous and the 
other for alcoholic solution. Each should be used with its intended 
solvent. Make a 1 per cent solution in alcohol or water. 

For material to be mounted whole in glycerin, glycerin jelly, or 
Venetian turpentine, stain several hours, or over night; pour off the 
stain, which may be used repeatedly; treat, without washing in 


58 


METHODS IN PLANT HISTOLOGY 


water, with a 2 per cent aqueous solution of acetic acid for 5 to 10 
minutes, changing the acid several times; transfer to 10 per cent 
glycerin, leaving about 1 c.c. of the acid to keep the whole solution 
slightly acid. When the glycerin becomes thick, mount in glycerin 
jelly. If the Venetian turpentine method is to be used, wash the glyc¬ 
erin out in alcohol slightly acidulated with acetic acid (a couple of 
drops of acetic acid to 50 c.c. of alcohol), and do not drain off the last 
alcohol too completely before transferring to the 10 per cent Venetian 
turpentine. According to Lee, the glycerin should be slightly alka¬ 
line. The alkalinity can be brought about by adding half a gram of 
common salt to 100 c.c. of the 10 per cent glycerin. We have found 
that eosin keeps better when the media are slightly acid. 

For staining paraffin sections, the alcoholic solution is better 
and the time may not be more than a few minutes, especially if the 
eosin is being used as a contrast stain. 

We have found the Eosin Y, of Coleman and Bell, very satis¬ 
factory, especially for fungi to be mounted whole. With the rapid 
improvement in the manufacture of stains, it is very probable that 
other dealers will have equally good products. Investigators will 
save time and money by keeping track of the findings of the commis¬ 
sion on Standardization of Biological Stains. 

Haematoxylin and eosin and methyl blue and eosin are good 
combinations. The eosin should follow the other stain. 

Erythrosin.—This is really an eosin, but there is some difference 
in the method of manufacturing. It is more precise and a more 
transparent stain than eosin and is to be preferred for nearly all 
staining of paraffin sections. Make a 1 per cent solution in distilled 
water or in 70 per cent alcohol. It gives good results when made up 
according to the general formula. 

Erythrosin stains rapidly, 30 seconds to 3 minutes being sufficient. 
When used in combination with other stains, erythrosin should come 
last. 

Magdala Red. —At least two Magdala reds are sold by dealers, 
one the echt (genuine) Magdala red, and the other simply Magdala 
red. The latter is much cheaper and, in our experience, much 
superior to the echt stain. The directions apply to the cheaper stain. 

For staining algae which are to be mounted in Venetian turpentine, 
use a 1 per cent solution in 85 or 95 per cent alcohol. Stain for 6 
to 8, or even 24, hours. Rinse in 95 and 100 per cent alcohol for a 


STAINS AND STAINING 


59 


few minutes. Transfer to 10 per cent Venetian turpentine and allow 
the turpentine to concentrate as described in chapter viii. 

In staining sections to be mounted in balsam, the same stain may 
be used, but it is better to dilute it one-half with water. Stain for 
6 to 24 hours, dehydrate in 95 per cent and absolute alcohol, clear 
in clove oil, and mount in balsam. 

Magdala red stains lignified, suberized, and cutinized structures, 
and also chromosomes, centrosomes, nucleoli, and pyrenoids. It is 
likely to overstain, but the differentiation is easily secured by placing 
the finished mounts upon a white background in the direct sunlight. 
When the desired differentiation has been reached, it is better to 
avoid direct sunlight, although the mounts do not seem to fade in 
the ordinary light of a room. 

Except for special purposes, it is better to use this stain in com¬ 
bination with blue, green, or violet. 

Magdala red (not echt) has always been erratic in its behavior, 
and recent analyses of samples show that it is variable in composi¬ 
tion. A stain called phloxine, made by the National Anilin and 
Chemical Co., behaves like the best Magdala red and seems to give 
uniform results. 

Gentian-Violet.—This is one of the most important stains in the 
botanical laboratory. It may be made according to the general 
formula for anilin stains, but that solution does not keep well. A 
1 per cent solution in distilled water keeps indefinitely and seems to 
be as good as, if not better than, the anilin solution. Gentian- 
violet dissolves readily in clove oil and some prefer to use it this way, 
rather than in water or alcohol. 

With the aqueous or anilin-oil solutions, the following directions 
will enable the student to become acquainted with the behavior of 
the stain. Transfer to the stain from water and allow the stain to 
act for 1 to 30 minutes. The time depends upon the fixing and upon 
the structures to be stained. The brilliancy of the stain in achromatic 
structures may often be increased by leaving the slide from 2 to 5 
minutes in a 1 per cent aqueous solution of permanganate of potas¬ 
sium before applying the stain. The greatest objection to the 
aqueous and anilin-oil solutions of gentian-violet is that the stain 
washes out so rapidly in alcohols that it is impossible to run the 
slide up through the series. The usual practice is to dip the slide 
in water to remove most of the stain and thus avoid carrying it into 


60 


METHODS IN PLANT HISTOLOGY 


the alcohol: then transfer directly from water to 95 per cent alcohol, 
allowing the alcohol to act for only 2 or 3 seconds, then allow the 
absolute alcohol to act for 5 or 6 seconds, and then, while the stain 
is still coming out in streams, begin the treatment with clove oil. 
Holding the slide in one hand, pour on a few drops of clove oil, 
and immediately drain off in such a way as to carry off the alcohol. 
This clove oil should not be used again. Then flood the slide repeat¬ 
edly with clove oil, pouring the clove oil back into the bottle. A 
50-c.c. bottle of clove oil is large enough. About 100 mounts can 
be cleared with 50 c.c. of this oil. The clove oil is a solvent of 
gentian-violet, but it dissolves the stain from some structures more 
rapidly than from others; e.g., the stain may be completely removed 
from the chromosomes while it is still bright in the achromatic 
structures. As soon as the stain is just right, drain off the clove oil 
and leave the slide in xylol for a minute or two before mounting in 
balsam. This is a necessary step, because the continued action of 
clove oil would cause the preparation to fade. As may be inferred 
from what has preceded, alcohol would soon extract the stain, without 
any application of clove oil. The clove oil is used, not only because 
it extracts the stain more slowly, but because it dissolves the stain 
from some structures more rapidly than from others; e.g., the 
stain may be completely removed from the chromosomes while it 
is still bright in the achromatic structures, so that with safranin 
and gentian-violet one can get red chromosomes on a violet 
spindle. 

Some still use cedar oil to follow the clove oil. This stops the 
action of the clove oil, but the preparations harden slowly. 

Gentian-violet is an excellent stain for achromatic structures in 
all stages of development. Chromatin, in many of its stages, is 
also stained. In metaphase and anaphase one should be able to 
get red chromosomes and violet spindles with safranin and gentian- 
violet. If the chromosomes also persist in retaining the violet, 
shorten the stain in gentian-violet. Cilia stain well; starch grains 
stain deeply, chromatophores less deeply, and lignified walls may 
not stain at all. One should be able to get red lignified walls and 
violet cellulose walls with safranin and gentian-violet. 

Cyanin.—This stain is also called Quinolein blue and Chinolin 
blue. Dissolve 1 g. of cyanin in 100 c.c. of 95 per cent alcohol and 
add 100 c.c. of water. The cyanin would not dissolve in 50 per cent 


STAINS AND STAINING 


61 


alcohol. We have not found Grtibler’s cyanin at all satisfactory with 
the foregoing formula. With the general formula the Griibler’s 
cyanin will not dissolve. We use a cyanin prepared by H. A. Metz 
& Co., 122 Hudson Street, New York. This cyanin dissolves com¬ 
pletely when made up according to the general formula. It stains 
rapidly, 5 to 10 minutes usually being sufficient. Chromosomes take 
a deep blue, but the spindle is only slightly affected. Lignified 
structures stain blue, while cellulose walls are scarcely affected and 
the stain is easily washed out. 

Iodine Green.—Use a 1 per cent solution in 70 per cent alcohol. 
Stain for an hour, rinse in 70 per cent alcohol, dehydrate in 95 per 
cent alcohol and absolute alcohol, clear in xylol or clove oil, and 
mount in balsam. If the stain washes out too rapidly and does not 
give sufficient differentiation, stain longer, over night or even 24 hours. 

Lignified structures stain green, but, after proper washing, 
cellulose is scarcely affected. A bright green may be left in the 
chromosomes after all the stain has been washed out from the spindle. 

Acid fuchsin, erythrosin, and eosin are good contrast stains 
for mitotic figures. Acid fuchsin or Delafield’s haematoxylin are 
good for cellulose walls. 

Light Green (Licht Griiri ).—Light green is an acid stain, soluble 
in water, alcohol, or clove oil. It stains quickly and forms a sharp 
contrast with safranin or Magdala red. 

Stain in safranin and then, with little or no washing out, stain in 
a weak alcoholic solution of acid green (about 0.2 g. in 100 c.c. of 
95 per cent alcohol). From 20 seconds to about 1 minute may be 
sufficient. The green rapidly reduces the safranin, and consequently 
the staining must not be too prolonged. A successful preparation 
should show red chromosomes and green spindle. Lignified walls 
should be red and cellulose walls green. 

Malachite Green.—A 1 to 3 per cent aqueous solution is good for 
cellulose walls. The stain contrasts well with Congo red. 

Methyl Green.—A 1 per cent solution in water is good for staining 
lignified structures. Lee recommends that the solution be acidulated 
with acetic acid. This is not necessary for staining lignified mem¬ 
branes nor for staining chromosomes. Methyl green has long been 
a favorite stain for living tissues. It is more easily controlled than 
iodine green, especially in double staining to differentiate lignified 
and cellulose walls. 


62 


METHODS IN PLANT HISTOLOGY 


Acid Green.—Make a solution according to the general formula, 
or simply make a 1 per cent solution in water. This stains cellulose 
walls and achromatic structures, but scarcely affects lignified walls 
or chromosomes. 

Anilin Blue.—Strong alcoholic solutions are best for botanical 
work. Even though the dry stain may be intended for aqueous 
solution, make a 1 per cent solution in 85 or 95 per cent alcohol. 

This stain can be recommended for cellulose walls, achromatic 
structures of mitotic figures, for cilia, and it is particularly valuable 
for algae. Directions for using it with algae are given in chapter viii. 

Orange G.—Make a 1 per cent solution in water, in 95 per cent 
alcohol, or in clove oil. We prefer the solution in clove oil. 

The orange dissolves very slowly if put directly into clove oil. 
It is better to dissolve 3 or 4 grams of orange in 50 c.c. of absolute 
alcohol. In a well-corked bottle in the paraffin oven at 52° C., this 
much orange may go into solution. Then remove the cork and allow 
about half of the alcohol to evaporate. Pour on 200 c.c. or more 
of clove oil and let it stand for several hours. If any of the stain 
has not gone into solution, pour off the clear fluid, which is now ready 
for use, and pour some more clove oil on the residue, allowing the 
residue to go slowly into solution. In staining, we use a small bottle 
of the orange, pouring it on the slide and draining it back into the 
bottle. The absolute alcohol, carried into the clove oil in this way, 
does no damage, except that it dilutes the stain a little. 

Transfer to the aqueous stain from water; to the alcoholic stain 
from 85 per cent alcohol, since the stain is always applied as a second 
or third stain; use the solution in clove oil after the dehydration in 
absolute alcohol. Times are always short and are to be reckoned in 
seconds rather than in minutes. If the solution in clove oil has 
been used, the slide should be transferred to xylol before mounting 
in balsam. 

This is a plasma stain. It is distinctly a general rather than a 
selective stain, but is valuable as a background for other structures 
which have been stained violet or blue or green. It first came into 
prominence as the third member of the triple stain, safranin, gentian- 
violet, orange. 

Gold Orange.—This stain, which many incorrectly suppose to 
be the same as orange G, is much more readily soluble in clove oil 
and stains with much greater rapidity. 


STAINS AND STAINING 


63 


Bismarck Brown.—Use a 2 per cent solution in 70 per cent al¬ 
cohol. 

This is a good stain for cellulose walls, although it is not so precise 
as haematoxylin. Embryo sacs stained in one of the carmines are 
improved by 1 or 2 minutes’ staining in Bismarck brown. Material 
fixed in alcohol stains better than that which has been fixed in reagents 
containing chromic acid. A faint background of Bismarck brown is 
quite effective in staining sections containing bacteria. 

Nigrosin.—Make a 1 or 2 per cent solution in water. A few 
drops of this solution to a watch glass full of water stains filamentous 
algae or fungi in 1 to 3 hours. It keeps well in glycerin or Venetian 
turpentine. It also keeps well in balsam, but it is of little value in 
staining microtome sections. 

COMBINATION STAINS 

Sometimes preparations are stained with a single stain, selected 
to emphasize some particular feature, but in the great majority of 
cases two or more stains are used. In staining a vascular bundle, 
one stain may be selected which stains the xylem, but not the phloem, 
while another of a different color stains the phloem, but not the 
xylem, thus affording a sharp contrast. In staining mitotic figures, 
one stain may stain the chromosomes, while another of a different 
color may be used to stain the spindle. 

Success in double staining can be obtained only by noting the 
effect of each stain upon the various plant structures. 

Flemming’s Safranin, Gentian-Violet, Orange.—Safranin has 
long been a famous stain for mitosis. This triple combination was 
published in 1891, but its value in plant cytology was not thoroughly 
appreciated until five or six years later, when its application was 
developed to a high degree of perfection by various investigators of 
the Bonn (Germany) school. Three methods, which may be desig¬ 
nated as A, B, and C, will be described. 

A. According to Flemming, 1 stain 2 or 3 days in safranin (dissolve 
0.5 g. safranin in 50 c.c. absolute alcohol, and after 4 days add 10 c.c. 
distilled water); rinse quickly in water; stain 1 to 3 hours in a 
2 per cent aqueous solution of gentian-violet; wash quickly in water, 
and then stain 1 to 3 minutes in a 1 per cent aqueous solution of 
orange G. Transfer from the stain to absolute alcohol, clear in clove 
oil, and mount in balsam. 

1 Flemming’s original method is included only as interesting history. 


64 


METHODS IN PLANT HISTOLOGY 


B. The following formulas and method seem to be better for 
mitotic phenomena in plants: Make a 1 per cent solution of alcoholic 
safranin in absolute or 95 per cent alcohol, and after the safranin is 
completely dissolved, add an equal volume of a 1 per cent solution 
of aqueous safranin in water, thus making a 1 per cent solution of 
safranin in 50 per cent alcohol. Use a 1 per cent aqueous solution 
of gentian-violet, and a 1 per cent aqueous solution of orange G. 

Transfer paraffin sections to the stain from 95 per cent alcohol 
after the xylol or turpentine used in dissolving away the paraffin has 
been rinsed off. Stain 3 to 24 hours. If the period be too short, 
the washing out is so rapid that it is difficult to stop the differentiation 
at the proper point and, besides, the red is likely to be less brilliant. 
Rinse in 50 per cent alcohol until the stain is properly differentiated. 
Leave the slide in the 50 per cent alcohol until the stain is washed 
out from the spindle and cytoplasm, but stop the washing out before 
the chromosomes begin to lose their bright red color. If the washing 
out takes place too slowly, treat with slightly acidulated alcohol 
(one drop of HC1 to 50 c.c. of 50 per cent alcohol) for a few seconds. 
The acid must be removed by washing for 15 to 30 seconds in alcohol 
which has not been acidulated. 

Then dip the slide 5 or 6 times into water and stain in gentian- 
violet. The time required is so variable that definite instructions 
are impossible. The gentian-violet should stain the spindle, but 
not the chromosomes. If the stain be too prolonged, it may be 
impossible to get it out from the chromosomes and still leave it bright 
in the spindle. If the period be too short, the stain will wash out 
from the spindle. For mitotic figures in the germinating spores of the 
liverwort, Pellia, 30 minutes is not too long. In this case, the stain 
washes out easily from the chromosomes without the use of acid, 
and the spindle takes a rich violet which is not easily washed out. 
In embryo sacs of Lilium try 10 minutes. In pollen mother-cells try 
5 to 10 minutes. For root-tips try 2 to 10 minutes. Chromatin in 
the early prophases and in telophases will stain with the violet, and 
the violet will not wash out, but in phases in which fully formed 
chromosomes are visible the violet can be washed out if the period 
has not been too long. 

Remove the slide from the gentian-violet and dip it 5 or 6 times in 
water and then stain 30 seconds to 1 minute in orange G. The orange 
stains cytoplasm and at the same time washes out gentian-violet. 


STAINS AND STAINING 


65 


Transfer from the orange G to 95 per cent alcohol, dipping the 
slide a few times in this merely to save the absolute alcohol. Dehy¬ 
drate in absolute alcohol 3 to 30 seconds. 

Clear in clove oil, as already described in the paragraph on gentian- 
violet. Transfer to xylol and mount in balsam. 

Safranin and gentian-violet are often used without the orange. 
In this case, transfer from the gentian-violet directly to 95 per cent 
alcohol, and proceed as before. 

An objection to both these methods is that the gradual series 
of alcohols cannot be used, because the gentian-violet washes out so 
rapidly. If one should try a filament of Spirogyra with either of 
these methods, it would hardly be recognizable when it reached the 
balsam; but with thin sections, especially when well fastened to the 
slide, conditions are different and there does not seem to be any 
serious damage. 

Just now, we are using a third method, which seems better than 
either of the two just described. 

C. Use the safranin solution described in B, an aqueous solution of 
gentian-violet, and a solution of orange G or gold orange in clove oil. 

Transfer to the safranin from 50 per cent alcohol and stain as 
directed under B. Rinse in 50 per cent alcohol for a minute, then 
in water for a minute, and stain in gentian-violet 3 to 30 minutes. 
Rinse quickly in water, dehydrate for about 5 seconds in 95 per cent 
alcohol and for about 10 seconds in absolute alcohol; then pour on 
the slide the orange dissolved in clove oil and allow it to act for 10 
to 30 seconds. Drain off the clove-oil orange, which can be used 
again, pour on pure clove oil and watch until the gentian is satisfac¬ 
tory, then transfer to xylol and balsam. 

With freehand sections, which are likely to be much thicker, 
the process is the same but the times will be longer. 

Safranin and Light Green.—Stain in aqueous safranin; wash in 
water; pass through 50, 95, and 100 per cent alcohol, about 2 minutes 
in each grade; stain in light green dissolved in clove oil; rinse in xylol 
and mount in balsam. 

If the stain does not rinse off readily, transfer to absolute alcohol, 
then to equal parts xylol and absolute alcohol, clear in xylol and 
mount in balsam. 

Cyanin and Ery thro sin.—Both solutions may be made according 
to the general formula for anilins, or 1 per cent aqueous solutions 


66 


METHODS IN PLANT HISTOLOGY 


may be used. Miss Thomas recommends 1 gram dissolved in 100 
c.c. of 95 per cent alcohol. After the solution is complete, add 
100 c.c. of distilled water. Since the combination is used only for 
paraffin sections or for small organisms dried down on the slide, her 
formula is preferable. Transfer to the alcoholic cyanin from 50 per 
cent alcohol, stain 5 to 10 minutes or longer; rinse quickly in 50 
per cent alcohol, transfer to erythrosin and stain 30 seconds to one 
minute. Rinse quickly in 50 per cent alcohol, then in 95 per cent and 
absolute alcohol. Clear in xylol and mount in balsam. 

If aqueous stains are used, transfer to the cyanin from water, 
rinse in water, stain in erythrosin, rinse in water, and transfer directly 
to 95 per cent alcohol. If the cyanin washes out, stain for 1 hour, 
and if it still washes out, omit the rinsing and transfer directly from 
the cyanin to the erythrosin. 

The erythrosin may be used first; in this case stain for 5 minutes 
in erythrosin, transfer directly to cyanin, and stain for about 10 
seconds. Dehydrate in 95 per cent and in absolute alcohol, clear 
in xylol or in clove oil, and mount in balsam. 

The stains wash out so rapidly that the series of alcohols cannot 
be used. 

Chromosomes and nucleoli stain blue and achromatic structures 
red. Lignified structures stain blue and cellulose walls red. The 
various cell constituents are often sharply differentiated. It was this 
combination which suggested the now obsolete terms, “cyanophilous” 
and “erythrophilous.” 

Magdala Red and Anilin Blue.—Make both solutions as directed 
in chapter viii on “The Venetian Turpentine Method.” 

For paraffin sections, stain 3 to 24 hours in Magdala red, dip in 
95 per cent alcohol to rinse off the stain, and then stain 2 to 10 minutes 
in the anilin blue. Dip in 95 per cent alcohol to rinse off the stain, 
and treat for a few seconds with alcohol slightly acidulated with 
hydrochloric acid (one drop to 50 c.c. of 95 per cent alcohol). In the 
acid alcohol the blue will become more intense, but the red would 
soon be extracted. Wash in 95 per cent alcohol to remove the acid. 
If the acid has weakened the Magdala red, put a pinch of sodium 
carbonate into the 95 per cent alcohol. The red may brighten. If 
the red is too weak, return to the Magdala red and try again. From 
the 95 per cent alcohol, transfer to absolute alcohol, to xylol, and then 
mount in balsam. 


STAINS AND STAINING 


67 


Acid Fuchsin and Iodine Green Mixtures.—Two solutions are 
kept separate, since they do not retain their efficiency long after 


they are mixed: 

^ f Fuchsin acid. 0.1 g. 

\ Distilled water. 50.0 c.c. 

g f Iodine green. 0.1 g. 

\ Distilled water. 50.0 c.c. 

{ Absolute alcohol. 100.0 c.c. 

Glacial acetic acid. 1.0 c.c. 

Iodine. 0.1 g. 


Mix equal parts of A and B. Transfer to the stain from water. 
The proper time must be determined by experiment. For a trial, 
24 hours might be recommended. Transfer from the stain directly 
to solution C and from C to xylol. 


A. Acid fuchsin. 0.5 g. 

B. Iodine green. 0.5 g. 


Mix a pipette full of A with a pipette full of B; stain 2 to 8 
minutes; transfer to 85 per cent or 95 per cent alcohol, dehydrate 
rapidly, clear in xylol, and mount in balsam. Both these formulas 
are good for mitosis. 

Acid Fuchsin and Methyl Green.—Both may be used in 1 per 
cent aqueous solutions. 

For mitotic figures, stain in green for about an hour, wash in 
water or alcohol until the green is extracted from the spindle, and 
then stain for about 1 minute in the fuchsin. Dehydrate in 95 
and 100 per cent alcohol, clear in xylol or clove oil, and mount in 
balsam. If the green washes out, stain longer; if it is not readily 
extracted from the spindle, shorten the period. If the fuchsin stains 
the chromosomes, shorten the period, and lengthen it if the fuchsin 
washes out from the spindle. The chromosomes should take a 
brilliant green and the spindle a bright red. 

Delafield’s Haematoxylin and Erythrosin.—Stain first in the 
haematoxylin, and after that stain is satisfactory, stain for 30 seconds 
or 1 minute in erythrosin. This is a good combination, and, for 
most plant structures, gives a far better differentiation than the 
traditional haematoxylin and eosin, since the erythrosin has all the 
advantages of the eosin and is more transparent. Orange G is also 
a good stain to use with Delafield’s haematoxylin. 











68 


METHODS IN PLANT HISTOLOGY 


Directions for staining in safranin and Delafield’s haematoxylin 
are given in the chapter on “ Freehand Sections” (chap. vi). 

Haidenhain’s Iron-Haematoxylin and Orange G.—This haema¬ 
toxylin is very satisfactory when used alone. A light staining in 
orange G, however, sometimes improves the mount. After the last 
washing in water, stain for about 30 seconds in orange G, or, if the 
orange is in clove oil, stain after dehydrating in absolute alcohol. 
Eosin, erythrosin, and nearly all plasma stains fail to increase the 
effect of a good stain in iron-haematoxylin. 

Combinations might be described almost without limit. Several 
more will be suggested in connection with the various groups of plants 
in Part II. 

We have not attempted to make the list of stains complete. It is 
better to master a few stains than to use many stains indifferently. 
A successful photographer once advised an amateur to stick to one 
brand of plate and one formula for developer. His hint might well 
have a wider application. If one really masters two or three good 
combinations, he is well prepared to develop methods for meeting 
special needs. 


CHAPTER IV 

GENERAL REMARKS ON STAINING 

Many things may be examined alive without killing, fixing, 
staining, or any of those processes. A filament of Spirogyra shows 
the chromatophore nicely if merely mounted in a drop of water; 
the nucleus may be visible and the pyrenoids can usually be located. 
Of course, such a study is necessary if one is to understand anything 
about the plant, and in an elementary class this might be sufficient; 
but a drop of iodine solution applied to the edge of the cover would 
emphasize certain details, e.g., the starch would appear blue, the 
nucleus a light brown, and the cytoplasm a lighter brown. This 
illustrates at least one advantage to be gained by staining; it enables 
us to see structures which would otherwise be invisible, or almost 
invisible. Much of the recent progress in morphology and cytology 
has been due to the development of critical methods of staining. 
Some of the combinations and methods recommended by various 
workers are good in themselves, while others, not so good, have 
yielded results because they have been so skilfully used. 

SELECTION OF A STAIN 

With so many stains at our disposal, it at once becomes a problem 
just which stain or combination to use in each particular case. 
Beautiful and instructive preparations occasionally result from some 
happy chance, but uniform success demands skill and judgment in 
manipulation, and also a knowledge of the structures which are to 
be differentiated. Let us take a vascular bundle for illustration. 
Safranin stains the xylem a bright red, but, with judicious washing, 
is entirely removed from the cambium and cellulose elements of the 
phloem. A careful staining with Delafield’s haematoxylin now gives 
a rich purple color to the cellulose elements which were left unstained 
by the safranin, thus contrasting sharply with the lignified elements. 
If cyanin and erythrosin be used, the xylem takes the blue while the 
cambium and phloem take the red. 

The mere selection of two colors which contrast well is not 
sufficient. Green and red contrast well, but safranin and iodine 

69 


70 


METHODS IN PLANT HISTOLOGY 


green would be a poor combination, for both would stain chromo¬ 
somes and neither would stain the spindle; both would stain lignified 
structures and neither would give satisfactory results with cellulose 
walls. Both stains are basic. Acid green would have given a con¬ 
trast in both these cases, because it stains achromatic structures and 
cellulose walls. In general, an acid stain should be combined with 
a basic one, but there are so many exceptions that it is hardly 
worth while to learn a list of basic and acid stains. Stains which 
stain chromosomes are likely to be basic, and those which do not stain 
chromosomes are likely to be acid or neutral. If it were true that 
acid stains affect only basic structures, and basic stains affect only 
acid structures, a classification of stains would be of great value. 
Safranin and gentian-violet are both basic, but with proper washing 
out the chromosomes are red and the spindle is violet, the safranin 
being washed out from the spindle, while the gentian-violet is washed 
out from the chromosomes. The only way to insure success is to 
become familiar with the action of each stain upon the various 
structures. 

THEORIES OF STAINING 

In 1890 Auerbach, a zoologist, published the results of his studies 
upon spermatozoa and ova. He found that, if preparations contain¬ 
ing both spermatozoa and ova were stained with cyanin and erythrosin, 
the nuclei of the spermatozoa took the cyanin, while the nuclei of 
ova preferred the erythrosin; hence he proposed the terms “cyano- 
philous” and “erythrophilous.” Auerbach regarded these differ¬ 
ences as an indication of sexual differences in the cells. 

Rosen (1892) supported this theory, and even went so far as to 
regard the tube nucleus of the pollen grain as female, on account of its 
erythrophilous staining. In connection with this theory it was 
suggested that the ordinary vegetative nuclei are hermaphrodite, 
and that in the formation of a female germ nucleus the male elements 
are extruded, leaving only the erythrophilous female elements; 
and, similarly, in the formation of a male nucleus the female 
elements are extruded, leaving only the cyanophilous male elements. 

As long ago as 1884 Strasburger discovered that with a mixture 
of fuchsin and iodine green the generative nucleus of a pollen grain 
stains green, while the tube nucleus stains red. In 1892, in his 
Verhalten des Pollens, he discussed quite thoroughly the staining 
reactions of the nuclei. The nuclei of the small prothallial cells of 


GENERAL REMARKS ON STAINING 


71 


gymnosperm microspores are cyanophilous like the male generative 
nuclei. The nuclei of a nucellus surrounding an embryo sac are also 
cyanophilous, while the nuclei of structures within the sac are erythro- 
philous. His conclusion is that the cyanophilous condition in both 
cases is due to poor nutrition, while the erythrophilous condition is 
due to abundant nutrition. A further fact in support of the theory 
is that the nuclei of the adventitious embryos which come from the 
nucellus of Funkia ovata are decidedly erythrophilous, while the 
nuclei of the nucellus to which they owe their food-supply are 
cyanophilous. 

In division stages nuclei are cyanophilous, but from anaphase 
to resting stage the cyanophilous condition becomes less and less 
pronounced, and may even gradually change to the erythrophilous. 

An additional fact in favor of this theory is that in Ephedra the 
tube nucleus, which has very little cytoplasm about it, is cyanophilous. 
Strasburger claimed that there is no essential difference between 
male and female generative nuclei, and subsequent observation soon 
showed that within the oospore the sex nuclei rapidly become alike 
in their reaction to stains. 

Malfatti (1891) and Lilienfeld (1892-1893) claim that these 
reactions are dependent upon the amount of nucleic acid present in the 
structures. During mitosis the chromosomes consist of nearly 
pure nucleic acid and are intensely cyanophilous, but the proto¬ 
plasm, which has little or no nucleic acid, is erythrophilous. There is 
a gradual transition from the cyanophilous condition to the erythro¬ 
philous, and vice versa, the acid structures taking basic stains and 
basic structures the acid stains. 

The terms “erythrophilous” and “cyanophilous” soon became 
obsolete, and many claimed the affinity is for basic and acid dyes, 
rather than for blue or red colors. That the terms were misnomers 
became evident when a combination like safranin (basic) and acid 
green (acid) was used, for the cyanophilous structures stained red, 
and the erythrophilous green. 

According to Fischer (1897 and 1900), stains indicate physical 
but not chemical composition. Fischer experimented with sub¬ 
stances of known chemical composition. Egg albumin was shaken 
until small granules were secured. These were fixed with the usual 
fixing agents, and then stained with Delafield’s haematoxylin. The 
extremely small granules stained red, while the larger ones became 


72 


METHODS IN PLANT HISTOLOGY 


purple. Since the granules are all alike in chemical composition, 
Fischer concluded that the difference in staining must be due to 
physical differences. With safranin, followed by gentian-violet, 
the larger granules stain red and the smaller violet; if, however, the 
gentian-violet be used first, then treated with acid alcohol and fol¬ 
lowed by safranin, the larger granules take the gentian-violet and 
the smaller the safranin. In root-tips similar results were obtained. 
Safranin followed by gentian-violet stained chromosomes red and 
spindle fibers violet, while gentian-violet followed by safranin stained 
the chromosomes violet and the spindle red. One often reads that 
chromosomes owe their strong staining capacity to nuclein, and 
especially to the phosphorous, but, according to Fischer, this is 
shown to be unfounded, since albumin gives similar results, yet 
contains no phosphorous, and is not chemically allied to nuclein. 

Probably the most important reason which led Fischer to under¬ 
take this series of experiments was the claim that certain granules 
of the Cyanophyceae should be identified as chromatin because they 
behaved like chromatin when stained with haematoxylin. Fischer’s 
experiments not only proved that chromatin cannot be identified 
in this way but raised the question whether staining reactions ever 
indicate chemical composition. At present, it would seem that, in 
most cases, the staining indicates only physical differences. How¬ 
ever, in some cases there is a chemical reaction, e.g., when material 
fixed in bichloride of mercury is stained in carmine, mercuric carminate 
is formed. 

It would be very convenient if we knew just how much depend¬ 
ence should be placed upon staining reactions as a means of analysis. 
If two structures stain alike with Delafield’s haematoxylin, does this 
mean that they have the same chemical composition; or if, on the 
other hand, they stain differently, must they necessarily be different 
in their chemical composition? Delafield’s haematoxylin, when 
carefully used, gives a rich purple color, but a careful examination 
will often show that in the same preparation some structures stain 
purple, while others stain red. Does this mean that the purple and 
red structures must have a different chemical composition ? Many 
people believe that structures which stain differently with a given 
stain must be chemically different, but they readily agree that struc¬ 
tures which stain alike are not necessarily similar in chemical com¬ 
position. Chromosomes of dividing nuclei and lignified cell walls 


GENERAL REMARKS ON STAINING 


73 


stain alike with safranin; chromosomes and cellulose cell walls stain 
much alike with Delafield’s haematoxylin; but everyone recognizes 
that the chromosome is very different in its chemical composition 
from either the cellulose or the lignified wall. 

However, in an indirect and somewhat uncertain way, one can 
infer the nature of certain structures from the staining. For instance, 
if sections of various objects have been stained with safranin, we may 
draw the following inferences with more or less confidence: if cells 
in the xylem region of a vascular bundle stain red, their walls are 
lignified; if cortical cells, which may appear quite similar in transverse 
section, stain red, they are likely to be suberized; if the outer walls 
of epidermal cells stain red, they are cutinized; but if the outer 
boundary of the embryo sac of a gymnosperm stains red, it is chitin- 
ized. Of course, these inferences can be made only because the 
various structures have been tested by more accurate methods. 

Whatever doubt or uncertainty there may be in regard to theories 
of staining or in regard to the value of stains as a means of analysis, 
there is no doubt that stains are of the highest importance in differ¬ 
entiating structures, and in bringing out details which would other¬ 
wise be invisible. 


PRACTICAL HINTS ON STAINING 

The number of stains in the catalogues is becoming so great that 
it is impossible to become proficient in the use of all of them. As 
we have already intimated, it is better to master a few of the most 
valuable stains than to do indifferent work with many. An experi¬ 
enced technician knows that it is impossible to judge from a few 
trials whether a given stain or combination is really valuable or 
not. As a matter of fact, some of the most valuable combinations, 
like Haidenhain’s iron-alum haematoxylin and Flemming’s safranin, 
gentian-violet, orange, require patient study and long practice before 
they yield the magnificent preparations of the trained cytologist. 
The beginner, especially if somewhat unacquainted with the details 
of plant structure, may believe that he has an excellent preparation 
when it is really a bad, or at most an indifferent, one. To illustrate, 
let us suppose that sections of the pollen grain of a lily have been 
stained in safranin and gentian-violet. If the preparation merely 
shows a couple of dense nuclei and a mass of uniform cell contents 
surrounded by a heavy wall, the mount is poor. If the two nuclei 


74 


METHODS IN PLANT HISTOLOGY 


are quite different and starch grains are well differentiated in the 
tube cells and the wall shows a violet intine contrasting sharply 
with a red exine, the mount is good. Anything intermediate is 
indifferent. If mitotic figures have been stained with cyanin and 
erythrosin, a first-class preparation should show blue chromosomes 
and red spindles; if stained with safranin and gentian-violet, the 
chromosomes should be red and the spindles violet. 

In staining growing points, apical cells, young embryos, anther- 
idia, archegonia, and many such things, the cell walls are the principal 
things to be differentiated, if the preparations are for morphological 
study. As a rule, it is better in such cases not to use double stain¬ 
ing, but to select a stain which stains the cell walls deeply without 
obscuring them by staining starch, chlorophyll, and other cell con¬ 
tents. For example, try the growing point of Equisetum. The 
protoplasm of such growing points is very dense. If Delafield’s 
haematoxylin and erythrosin be used, the haematoxylin will stain 
the walls and nuclei, and will slightly affect the other cell contents, 
but the erythrosin will give the cytoplasm such a dense stain that 
the cell walls will be seriously obscured. It would be better to use 
haematoxylin alone. For counting chromosomes, it is better to stain 
in iron-alum haematoxylin alone, or in safranin alone. The same 
suggestion may well be observed in tracing the development of 
antheridia, archegonia, embryos, and similar structures. 

In using combinations, it must be remembered that the second 
stain often affects the first, e.g., if safranin is to be followed by 
Delafield’s haematoxylin in staining a vascular bundle, it will not 
do to make the safranin just right and then apply the haematoxylin, 
for the acid which must be used to differentiate the haematoxylin 
and to avoid precipitates will also reduce the safranin, and the red 
will be too weak. You must overstain in safranin so that the reduc¬ 
tion will finally leave it just right. The same hint will apply if 
safranin is to be followed by anilin blue, since here, also, acid must be 
used; if light green is to follow the safranin, the stain itself is so 
acid that the safranin must be rather strong before the light green is 
applied. Orange, whether in water or in clove oil, reduces many 
stains and, consequently, such stains must be strong enough to allow 
the weakening. These hints are only samples: the student must 
observe the behavior of the various stains when used singly and when 
used in various combinations. 


GENERAL REMARKS ON STAINING 


75 


The best advice that can be given, not only to the beginner but 
to the investigator is this: Master a few stains. For nuclear phe¬ 
nomena, strive for perfection in the use of Haidenhain’s iron-alum 
haematoxylin; then, if you can afford the time, practice the safranin, 
gentian-violet, orange combination. For vascular anatomy, learn to 
stain xylem with safranin; and, for a contrast, stain cellulose walls with 
Delafield’s haematoxylin, gentian-violet, light green or anilin blue. 
For filamentous algae and fungi, master the iron-alum haematoxylin 
method; and then try the Magdala-red and anilin-blue combination. 
Do not pass judgment against a standard method or even a new 
method just because you fail to get results at the first trial. After 
you have become proficient with the iron-alum haematoxylin for 
mitotic figures, you are sure to fail if you try the same procedure with 
Rhizopus; but, nevertheless, the stain is just as good for Rhizopus 
as for figures in pollen mother-cells or root-tips. 

Permanent preparations are an absolute necessity for the greater 
part of most advanced work, but let us not imagine that we cannot 
examine anything until we have made a permanent mount. It 
would be impossible to make a permanent mount of the rotation of 
protoplasm. It is better for many purposes to look at motile spores 
while they are moving. Use Spirogyra while it is fresh and green, 
and use permanent preparations only to bring out nuclei and other 
details which are not so easily seen in living material. Examples 
might be multiplied. 


CHAPTER V 

TEMPORARY MOUNTS AND MICROCHEMICAL TESTS 

Skill in making freehand sections, without any microtome, and 
in teasing with needles and in making delicate dissections under 
the simple microscope are absolutely necessary in any investigation 
dealing with the structure and development of plants. Preliminary 
study with the aid of such methods not only gives a broader view of 
structures in all dimensions and helps the interpretation of stained 
microtome preparations, but is necessary in determining whether 
material is worth all the labor of making permanent mounts. That 
particular class of temporary mounts intended only for chemical 
tests is considered separately in the second part of this chapter. 

TEMPORARY MOUNTS 

A preliminary examination of almost any botanical material 
may be made without any fixing, imbedding, or staining. If a little 
starch be scraped from a potato, and a small drop of water and a 
cover-glass be added, a very good view will be obtained, and if a 
small drop of iodine solution be allowed to run under the cover, the 
preparation, while it lasts, is better than some permanent mounts. 
The unicellular and filamentous algae can be studied quite satis¬ 
factorily from such mounts. The protonema of mosses and the 
prothallia of ferns should be studied in this way, even if a later 
study from sections is intended. The addition of a little iodine 
identifies the starch and makes the nucleus more plainly visible. 
If the top of a moss capsule be cut off at the level of the annulus, a 
beautiful view of the peristome may be obtained by simply mounting 
in a drop of water, or, in a case like this where no collapse is to be 
anticipated, the object may be mounted in a small drop of glycerin— 
just enough to come to the edge of the cover without oozing out 
beyond—and the preparation may be made permanent by sealing 
with balsam, gold size, or any good cement. The antheridia and 
archegonia of mosses may be examined if the surrounding leaves are 
carefully teased away with needles. Freehand sectioning with a sharp 

76 


TEMPORARY MOUNTS AND MICROCHEMICAL TESTS 77 

razor and judicious teasing with a pair of needles will give a fair insight 
into the anatomy of the higher plants without demanding any further 
knowledge of technic. This rough work is a very desirable ante¬ 
cedent to the study of microtome sections, because most students see 
in a series of microtome sections only a series of sections when, in the 
mind’s eye, they ought to see the object building itself up in length, 
breadth, and thickness as they pass from one section to another. 

The movements of protoplasm can, of course, be studied only in 
the living material. Every laboratory should keep Chara growing 
at every season of the year. Mount a small portion and note the 
movements in the internodal cells. Avoid any pressure and any 
lowering of the temperature. A gentle raising of the temperature 
will accelerate the movements. A leaf of Elodea shows the move¬ 
ments very clearly, especially in the midrib region. The stamen hairs 
of Tradescantia have long been used, their color, resembling a faint 
haematoxylin stain, making them particularly favorable. Stinging 
hairs show a brisk movement if they are mounted quickly and without 
injury. Fortunately, the common onion always furnishes favorable 
material for demonstrating the movements of protoplasm. Strip 
the epidermis from one of the inner scales of the bulb and mount 
in water. The granules may appear to better advantage in yellow 
light, like that of an ordinary kerosene lamp. 

The discharge of spores and gametes should be observed in the 
living material: the difference in the behavior of spores and gametes 
is very striking and can be appreciated only while they are alive. 
Most aspects of growth and movement can be studied best in the 
living condition. In short, it is well to make a preliminary study of 
everything. The germination of spores and the growth of pollen 

tubes can be studied in _ 

the hanging drop. For , LI.. 1 1 . .. [ 

facilitating such cultures ^ 17 _ The hanging ^ rop culture 

there are many devices, 

such as hollow-ground slides, glass rings, rubber rings, etc. (Fig. 17). 
A device which is better for most purposes, and which is easily made 
by any student, is shown in Figure 18. 

A square or round hole f inch in diameter is cut in a piece of 
pasteboard | inch thick, 1 inch wide, and 1J inches long. The paste¬ 
board is then boiled to sterilize it and to make it fit more closely to 
the slide. While the pasteboard is still wet, press it to the slide, make 





78 


METHODS IN PLANT HISTOLOGY 


the culture in a drop of water or culture solution on the cover, and 
invert the cover over the hole. A little.water added at the edge 
of the pasteboard from time to time will'keep it from warping and 
will at the same time provide a constant moist chamber. 



Fig. 18.—Another hanging-drop culture 


In collecting material for mitotic figures in anthers it is necessary 
to examine fresh anthers, if one wishes to avoid a tedious and uncer¬ 
tain search after the anthers have been imbedded. By teasing 
out a few cells from the apex and a few from the base of the anther 
the stage of development is readily determined, and anthers which 
do not show the desired stages can be rejected. By allowing a drop 
of eosin or methyl green to run under the cover the figures are more 
easily detected. The actual progress of mitosis has been observed 
in living stamen hairs of Tradescantia. 

MICROCHEMICAL TESTS 

Botanical microchemistry has developed to such an extent that 
it has become an independent subject, like bacteriology. We shall 
consider only the commonest tests which are needed constantly by 
students of morphology. For a thorough presentation of the chem¬ 
istry of the cell, we are looking forward with great anticipation to a 
forthcoming book by Dr. Sophia Eckerson, whose critical tests and 
analyses we have observed for many years. In the meantime, 
Pflanzenmikrochemie , by Dr. O. Tunmann (Gebriider Borntraeger, 
Berlin), is recommended to those who read German. Zimmerman’s 
Botanical Microtechnique (Henry Holt & Co., New York) is still 
recommended to those who must rely upon English. We shall give 
only a few tests, but in considering the various stains we shall indicate 
the effect of each stain upon the various plant structures. 

Starch.—Mount the starch or starch-containing structures in 
water, and allow a drop of iodine solution to run under the cover. 
Starch assumes a characteristic blue color. The solution may be 
prepared by dissolving 1 g. of potassium iodide in 100 c.c. of water 
and adding 0.3 g. of sublimed iodine. A strong solution of iodine in 
alcohol (about 1 g. in 50 c.c. of absolute alcohol) keeps well. A drop 






TEMPORARY MOUNTS AND MICROCHEMICAL TESTS 79 


of this solution added to 1 c.c. of water is good for testing. With too 
strong a solution, the starch first turns blue but rapidly becomes black. 

Grape-Sugar.—In cells containing grape-sugar, bright-red gran¬ 
ules of cuprous oxide are precipitated by Fehling’s solution. It is 
better to keep the three ingredients in separate bottles, because the 


solution does not keep long after they are mixed. The solutions 
may be labeled A, B, and C. 

^ f Cupric sulphate. 3 g. 

\ Water. 100 c.c. 

g f Sodium potassium tartrate (Rochelle salt).. 16 g. 

p f Caustic soda. 12 g. 

\ Water. 100 c.c. 


When needed for use, add to 10 c.c. of water 5 c.c. from each of 
the three solutions. The sections, which should be two or three 
cells in thickness, are warmed in the solution until little bubbles are 
formed. Too much heat must be avoided. Mount and examine in 
a few drops of the solution. The twig or organ may be treated with 
the solution, and the sections may be cut afterward. Other sub¬ 
stances precipitate copper, and may be mistaken for grape-sugar 
by the beginner. 

Cane-Sugar.—Cuprous oxide is not precipitated from Fehling’s 
solution by cane-sugar, but after continued boiling in this solution 
the cane-sugar is changed to invert-sugar and the copper is precipi¬ 
tated. The solution becomes blue. 

Proteids.—The proteids turn yellow or brown with the iodine 
solution. It is better to use a stronger solution than when testing 
for starch. It must be remembered that many other substances 
also turn brown when treated with iodine. 

When proteids are warmed gently in concentrated nitric acid, 
the acid becomes yellow. The color may be deepened by the addi¬ 
tion of a little ammonia or caustic potash. 

When proteids are heated with Millon’s reagent, the solution 
becomes brick-red or rose-red. This reaction takes place slowly 
even in the cold. The following is one formula for this reagent: 


Mercury./. 1 c.c. 

Concentrated nitric acid. 9 c.c. 

Water. 10 c.c. 


Dissolve the mercury in the nitric acid and add the water. 










80 


METHODS IN PLANT HISTOLOGY 


Fats and Oils.—The fatty oils are not soluble in water and are 
only slightly soluble in ordinary alcohol. They dissolve readily in 
chloroform, ether, carbon disulphide, or methyl alcohol. 

Alcannin colors oils and fats deep red. The test is not decisive, 
because ethereal oils and resins take the same red color. Dissolve 
commercial alcannin in absolute alcohol, add an equal volume of 
water, and filter. The fats and oils in sections left in this solution 
for 24 hours should be bright red. The reaction is hastened by 
gentle heating. 

Osmic acid, as used in fixing agents, colors fats and oils brown 
or black. The dark color is removed by bleaching in a 3 to 10 per cent 
solution of hydrogen peroxide. 

In case of fats and oils, solubility and color reactions are useful, 
but must be regarded as corroborative evidence, not as decisive 
proof. For more critical and detailed methods, consult the book by 
Tunmann, which will also give the literature of the subject. 

The Middle Lamella.—Even the origin and development of the 
middle lamella are none too well known; its microchemistry has 
progressed but little beyond the color-reaction stage. The middle 
lamella consists largely of pectin or pectic compounds. The easy iso¬ 
lation of cells, when treated with Schultze’s maceration, depends upon 
the ready solubility of pectins in this reagent. Many intercellular 
spaces arise through the natural solution or gelatinization of the lamella. 

In polarized light, with crossed Nichols, the middle lamella is 
resolved into three lamellae, the middle one appearing dark, and the 
two outer lamellae, light. 

Ruthenium red is a good stain, since it gives as good results as 
any and has the advantage of keeping well in balsam or glycerin 
jelly. Make a very weak solution—1 g. to 5,000 c.c. of water, 
or even weaker—and keep it in the dark. It stains many other 
things besides the lamella, but is, nevertheless, a good stain. 

Pectin is not at all confined to the middle lamella, but is found 
in other membranes, particularly in spore coats. 

Cellulose.—In concentrated sulphuric acid cellulose swells and 
finally dissolves. It is also soluble in cuprammonia. The cupram- 
monia can be prepared by pouring 15 per cent ammonia water upon 
copper turnings or filings. Let the solution stand in an open bottle. 
It does not keep well, but its efficiency is readily tested. Cotton 
dissolves almost immediately as long as the solution is fit for use. 


TEMPORARY MOUNTS AND MICROCHEMICAL TESTS 81 


With iodine and sulphuric acid cellulose turns blue. Treat 
first with the undiluted iodine-potassium-iodide solution described 
in the test for starch, then add a mixture of two parts of concentrated 
sulphuric acid and one part of water. 

With chloroiodide of zinc cellulose turns violet. Dissolve com¬ 
mercial chloroiodide of zinc in about its own weight of water and add 
enough metallic iodine to give the solution a deep-brown color. 

The cell walls of fungi consist of fungus cellulose. When young, 
they give a typical cellulose reaction; when older, they become 
insoluble in cuprammonia and, with iodine and sulphuric acid, show 
only a yellow or brown, instead of the typical blue. With chloroiodide 
of zinc, the wall stains yellow or brown, instead of violet. 

Reserve cellulose, which is common in thick-walled endosperm of 
seeds, shows the same microchemical reactions as ordinary cellulose. 

Callose.—The thickening on the sieve plate differs from cellulose 
in its staining reactions, and in its solubility. It is insoluble in 
cuprammonia, but will dissolve in a 1 per cent solution of caustic soda. 

Stain in a 4 per cent aqueous solution of soda (Na x CCh) for 
10 minutes, and transfer to glycerin. The callus should take a bright 
red. If stained very deeply and then transferred to a 4 per cent soda 
(without the corallin), the stain is extracted from the cellulose but 
remains in the callus. Unfortunately, the preparations are not 
permanent. 

If stained for about an hour in a dilute aqueous solution of anilin 
blue, the stain may be extracted with glycerin until it remains only 
in the callus. After the blue is satisfactory, a few minutes in aqueous 
eosin will afford a good contrast. The preparation may be mounted 
in balsam and is fairly permanent. 

Lignin.—Lignified walls are insoluble in cuprammonia. The 
iodine and sulphuric acid or the chloroiodide of zinc, used as in testing 
for cellulose, give the lignified walls a yellow or brown color. After 
a treatment with Schultze’s maceration fluid, lignified membranes 
react like cellulose. 

Phloroglucin in a 5 per cent aqueous or alcoholic solution applied 
simultaneously with hydrochloric acid gives lignified walls a reddish- 
violet color. The preparations do not keep. 

Cutinized and Suberized Walls.—These are insoluble in cupram¬ 
monia or concentrated sulphuric acid. They are colored yellow or 
brown by chloroiodide of zinc, or by iodine and sulphuric acid, when 


82 


METHODS IN PLANT HISTOLOGY 


applied as in testing for cellulose or lignin. With alcannin, they 
take a red color, but the red is not as deep as in case of fats and oils. 
After soaking in an aqueous solution of caustic potash, suberized 
membranes take a red-violet color when treated with chloroiodide 
of zinc. 

If a strong, fresh alcoholic solution of chlorophyll be allowed to 
act upon suberized membranes for 15 to 30 minutes in the dark, 
they stain green, while lignified and cellulose walls do not take the 
stain. The preparations are not permanent. 

A solution of alcannin in 50 per cent alcohol stains suberized and 
cutinized walls red, but the color may not be very sharp. 

Cyanin can be recommended. First, treat with Eau de Javelle 
(potassium hypochlorite), which can be obtained ready for use at 
any drug-store. This destroys tannins, and the lignified walls lose 
their staining capacity. Make a 1 per cent solution of cyanin 
(Griibler’s) in 50 per cent alcohol and add an equal volume of glycerin. 
This should show blue suberized walls, while the lignified walls 
remain unstained. 

Gum, Mucilage, and Gelatinized Membranes.—These are all 
soluble in water and are further characterized by their strong power 
of swelling. They are insoluble in alcohol. A series of forms with 
various color reactions is included under this heading. 

Crystals.—Nearly all crystals which are found in plants consist 
of calcium oxalate. Crystals of calcium carbonate, calcium tartrate, 
and calcium sulphate also occur. Calcium oxalate is soluble in 
hydrochloric acid or nitric acid. It is better to use the concentrated 
acids. The crystals are insoluble in water and acetic acid. Sulphuric 
acid changes calcium oxalate into calcium sulphate. When treated 
with barium chloride, crystals of calcium sulphate become covered 
with a granular layer of barium sulphate, while crystals of calcium 
oxalate are not affected. 

Calcium carbonate, when treated with hydrochloric acid or 
acetic acid, dissolves with effervescence. The acetic acid should be 
rather dilute. 


CHAPTER VI 


FREEHAND SECTIONS 

Sections which may be cut without imbedding, whether they are 
really cut freehand or with the aid of a microtome, will be considered 
here. The chapter will also deal with other small or thin objects 
which may be treated like freehand sections. 

The beginner is advised to start with the freehand section, because 
the processes are rapid, and it is comparatively easy to find the causes 
of imperfections and failures. In the paraffin method, where the 
processes are more complicated, it is often difficult, or even impossible, 
to determine the exact cause of a failure. 

As a matter of fact, real freehand sections, cut by holding the 
object in one hand and the knife in the other, are becoming less and 
less frequent in well-equipped laboratories. However, the laboratory 
is no place for one who is awkward with the hands; a certain amount 
of manual dexterity must be acquired if there is to be any success in 
morphological studies which demand critical preparations. Although 
we know the student will turn at once to the microtome, we venture 
a few remarks in regard to real freehand sections. 

A sharp razor is a necessity. For cutting sections of twigs, roots, 
rhizomes, and similar objects, a razor like the one shown in Figure 8 A, 
should be used; while for sections of soft tissues, like young aspara¬ 
gus stems, young ovaries of plants, most leaves, and such things, the 
type of razor shown in Figure 8 B, is much better. In cutting, brace 
the forearms against the sides, hold the object firmly in the left hand 
and cut with a long, oblique stroke from left to right. The edge of the 
razor and the direction of the stroke should be toward the body, 
not away from it as in whittling. If the material is fresh, the object 
and the razor should be kept wet with water, the razor being dipped 
in water for every stroke. For hard objects, like twigs of oak or 
maple, the razor will need sharpening after cutting a dozen sections. 
It is a waste of time to put off sharpening until the razor has become 
noticeably dull, for all sections except those cut when the razor is 
perfectly sharp are sure to be inferior. With softer material the 

83 


84 


METHODS IN PLANT HISTOLOGY 


razor may hold its edge for hundreds of sections. Those sections 
which seem to be worth further treatment should be placed at once 
in water or in a fixing agent and, of course, the choice of a fixing 
agent should be determined before the sections are cut. 

With the advent of a cheap, efficient sliding microtome, the hand 
microtome began to fall into disuse and, today, it has almost dis¬ 
appeared. 

The sliding microtome (Fig. 2) reduces to a minimum the necessity 
for manual dexterity, but it is a more complicated machine. Study 
the various parts before you begin to cut sections. How is the knife 
adjusted? How is the object clamp raised and lowered? How is 
the thickness of the section determined ? In case of a simple micro¬ 
tome like the one shown in Figure 2, the student should soon answer 
such questions without any help from the instructor. In case of 
more complicated microtomes, a demonstration by the instructor 
will save both time and machine. 

In cutting sections of wood or herbaceous stems, the knife should 
be set obliquely so as to use as much as possible of the cutting edge. 
In most cases it is neither necessary nor desirable to cut very thin 
sections by this method; 10 ju is very thin, and 20, 30, or even 40 n 
is usually thin enough. 

Cut with a firm, even stroke, wetting both knife and object after 
every section. Use water, if the material is fresh; if preserved, 
use the preservative. Some use a brush in removing sections from 
the knife, but nothing is quite equal to one’s finger; anyone who is 
in danger of a cut while performing this act is in need of this little 
practice in manual dexterity. 

WOODY AND HERBACEOUS SECTIONS 

Safranin and Delafield’s Haematoxylin.—In order to make the 
directions as explicit as possible, let us follow the processes from 
collecting the material to labeling the slide. The rhizome of Pteris 
aquilina is a good object to begin with. Dig down carefully until 
the rhizome is exposed; then with a sharp knife cut off pieces a few 
inches in length, taking the greatest care not to strain the tissues. 
If the rhizome has been cut carelessly or pulled up, as is usually the 
case, the finished mount will show ruptures between the bundles and 
bundle sheaths, disfiguring what might have been a beautiful prep¬ 
aration. 


FREEHAND SECTIONS 


85 


While the material is still fresh and moist, cut the sections and 
place them at once in 95 per cent alcohol, where they should remain 
20 to 30 minutes. It is not necessary to use a large quantity of 
alcohol; 10 c.c. is enough for 100 thin sections of the rhizome. 

Pour off the alcohol and pour on an alcoholic solution of safranin 
(a 1 per cent solution of safranin in 50 per cent alcohol. See chapter 
xxix on “Formulas for Reagents”). It is better to let the safranin 
act over night, or even for 24 hours. 

Pour off the safranin (which may be used repeatedly) and pour 
on 50 per cent alcohol. The alcohol will gradually wash out the 
safranin, but this stain is washed out more rapidly from cellulose 
walls than from those which are lignified. The sections should 
remain in the alcohol until the stain is nearly—but not quite—washed 
out from the cellulose walls, while still showing a brilliant red in the 
large lignified tracheids. If 5 or 10 minutes in the alcohol draws the 
safranin from the lignified walls as well as the cellulose, stain longer; 
if the differentiation is not secured in 5 or 10 minutes, a small drop of 
hydrochloric acid added to the alcohol will hasten the process. 
Some recommend staining for only 1 or 2 hours, but the washing-out 
process is likely to be rapid and uncertain. 

Pour off the alcohol and wash the sections thoroughly in ordinary 
drinking-water. The washing should be particularly thorough if 
acid has been used to hasten the previous process, for the preparations 
will fade if any acid remains. 

Stain in Delafield’s haematoxylin 3 to 30 minutes. Usually 5 
minutes will be about right. Delafield’s haematoxylin will stain the 
cellulose walls, but will have little or no effect upon lignified structures. 

Transfer to drinking-water, not distilled water. The red color 
of the whole section, as it appears to the naked eye, will be rapidly 
replaced by a rich purple. Continue to wash in water for 2 or 3 
minutes after the purple color appears. If the cellulose walls show 
only a faint purplish color, put the sections back into the stain and 
try a longer period. If the color is a deep purple or nearly black, 
add a little hydrochloric acid (1 drop to 50 c.c. is enough) to the 
water. It is better to put the drop into a bottle of water and shake 
thoroughly before letting the acidified water act upon the sections. 
As soon as the sections begin to appear reddish, which may be 
within 4 or 5 seconds, pour off the acidified water and wash in drinking- 
water, changing the water 3 or 4 times a minute, until the reddish 


METHODS IN PLANT HISTOLOGY 


color caused by the acid has been replaced by the rich purple color 
so characteristic of haematoxylin. The acid not only secures differ¬ 
entiation by dissolving out the stain from lignified structures more 
rapidly than from cellulose walls, but it also removes the disfiguring 
precipitates which almost invariably accompany staining with 
Delafield’s haematoxylin. The acid also washes out the safranin; 
it is for this reason that the washing after safranin should be stopped 
while there is still some red color in the cellulose walls. The acid 
should not only reduce the density of the haematoxylin and remove 
precipitates, but should also remove the little safranin which may 
remain in the cellulose walls. After the purple color has appeared, 
the sections should be left in water for 20 or 30 minutes. They 
might be left for several hours. 

Now place the sections in 50 per cent alcohol for 1 minute, then 
in 95 per cent alcohol for 1 minute, 100 per cent alcohol for 5 minutes, 
and then transfer to xylol. As soon as the sections become clear— 
in about 1 to 5 minutes—they are ready for mounting in balsam. If 
the sections do not clear readily, as may be the case if the air is 
damp, or if the alcohol or xylol is not quite pure, transfer from the 
absolute alcohol to clove oil, which will clear, even if the absolute 
alcohol is rather poor. Then transfer from clove oil to xylol; the 
objection to mounting directly from clove oil is that preparations 
harden more slowly than when mounted from xylol. With a section- 
lifter, or scalpel, or brush, transfer 3 or 4 sections to a clean, dry 
slide, put on 1 or 2 drops of balsam, and add a cover, first heating 
it gently to remove moisture. If xylol has been used for clearing, 
it is necessary to work rapidly; for the sections must never be allowed 
to dry. Use square or oblong covers for such mounts, reserving 
round covers for glycerin mounts. If material is abundant, use 
as many sections as you can cover conveniently. If you have used 
several stains with the same material, select for each mount sections 
from the different stains. In ordinary wood sections each mount 
should show the three most important views, transverse, longitudinal 
radial, and longitudinal tangential sections. It is wasteful to use 
three slides and three covers to show these three views, or to make a 
mount containing only a single section of the rhizome of Pteris. 

Put the label at the left. Write first the genus and species; then 
indicate what part of the plant has been mounted. The date on 
which the material was fixed is often valuable. After a year or so, 


FREEHAND SECTIONS 


87 


the date of making the mount may be of interest in indicating the 
relative durability of stains. The beginner is likely to write also the 
stains used, and other details, which he will find quite unnecessary 
after a little experience. Figure 19 illustrates a good style of labeling 
and mounting. 



Fig. 19.—The label 


The following is a convenient summary of the foregoing processes, 
beginning with the sections in 95 per cent alcohol: 

1. Sections in 95 per cent alcohol. 

2. Safranin, 12 to 24 hours. 

3. 50 per cent alcohol, with or without acid, until color is right, generally 
about 2 to 10 minutes. 

4. Water, 5 minutes, changing frequently. 

5. Delafield’s haematoxylin, 3 to 30 minutes. 

6. Water, 5 to 10 minutes, changing frequently. 

7. Water slightly acidulated, 5 to 10 seconds. 

8. Water, to wash out acid, 20 to 30 minutes. 

9. 50 per cent alcohol, 1 minute. 

10. 95 per cent alcohol, 1 minute. 

11. 100 per cent alcohol, 5 minutes. 

12. Xylol, 1 to 5 minutes. 

13. Balsam. 

14. Cover and label. 

If clove oil seems necessary, finish as follows: 

12. Clove oil, 2 to 5 minutes. 

13. Xylol, 1 to 5 minutes. 

14. Balsam. 

15. Cover and label. 

Another method has given excellent results, especially with old 
stems, pieces of dry boards, etc. Cut pieces 5 to 10 mm. square and 
2 to 3 cm. long, boil in water, and, when cool, transfer to equal parts 




















88 


METHODS IN PLANT HISTOLOGY 


of 95 per cent alcohol and glycerin. The material may be left in 
this mixture indefinitely. 

After sections have been cut, wash them in tap water, then in 
distilled water, and stain half an hour or more in weak Delafield’s 
haematoxylin—about 5 parts of the solution, as given in the formula, 
to 100 parts of water—and then wash in distilled water. Stain in 
weak safranin—about 2 parts of the stock solution to 100 parts of 
water—over night or even for several days. Wash in tap water, 
then wash in 95 per cent alcohol for 30 seconds or longer, according 
to the appearance of the stain. Dehydrate in absolute alcohol, 
clear in clove oil, transfer to xylol, and mount in balsam. This 
method is very good for gymnosperm woods. 

Since it usually happens that processes are commenced, but 
cannot be completed at a single laboratory period, it is necessary 
to know where sections may be left for several hours or until the 
next day without suffering injury. At 1, 2, or the pure water of 8 in 
the schedule given above, sections may be left until the next day. If 
it is not desirable to mount all of the sections which have been pre¬ 
pared, they may be kept indefinitely in clove oil or xylol. If the 
sections are to remain for a year or more in the clearing agent, xylol 
is to be preferred. Shells with good corks are best for keeping such 
material. 

For the study of vascular anatomy, this is the most permanent 
stain which has come into general use. 

More recently, safranin combined with anilin blue or with light 
green has been coming into favor. Both these methods will be 
described. 

Safranin and Anilin Blue.—Use the alcoholic safranin already 
described, and a 1 per cent solution of anilin blue in 90 per cent 
alcohol. 

With this combination we should recommend a long stain in saf¬ 
ranin, not less than 24 hours. Wash in 50 per cent alcohol, but do not 
extract all the safranin from the cellulose walls. Stain 2 to 10 minutes 
in anilin blue. Rinse a few seconds in 95 per cent alcohol, 
then treat for about 5 seconds with 95 per cent alcohol slightly 
acidulated with hydrochloric acid. The weak blue should at once 
change to a bright blue and, at the same time, the acid will remove 
some of the safranin. It is for this reason that we proceed while 
the sections are still somewhat overstained in safranin. Wash for 


FREEHAND SECTIONS 


89 


1 or 2 minutes in 95 per cent alcohol to remove the acid. A trace of 
sodium carbonate, just enough to make the alcohol alkaline, may 
be added to the 95 per cent alcohol. If any acid remains, the 
safranin will fade. Dehydrate in absolute alcohol 1 to 5 minutes, 
clear in xylol, or first in clove oil and then in xylol, and mount in 
balsam. 

For convenient reference, the process may be summarized, but 
it must be remembered that all the schedules are intended merely to 
introduce the method to the beginner. 

1. Sections in 95 per cent alcohol. 

2. Stain in safranin, 24 hours. 

3. 50 per cent alcohol until the stain becomes weak in cellulose walls, but 
not until it is removed entirely. 

4. Anilin blue, 2 to 10 minutes. 

5. 95 per cent alcohol, 2 to 5 seconds. 

6. 95 per cent alcohol, slightly acidulated with hydrochloric acid, 5 seconds. 

7. 95 per cent alcohol, with or without a trace of sodium carbonate, 1 or 2 
minutes. 

8. Absolute alcohol, 1 to 5 minutes. 

9. Xylol, 1 to 5 minutes. The xylol may be preceded by clove oil. 

10. Mount in balsam. 

Lignified and suberized walls should stain bright red and cellulose 
walls bright blue. To make this beautiful combination a success, it 
is necessary to be very careful. If too much safranin is extracted 
at stage 3, the acid at stage 6 will still further weaken the red stain 
and the contrast will not be sharp. 

Safranin and Light Green (Land’s Schedule).—This is another 
beautiful combination and the student should be successful from 
the first, since the light green is simpler to apply than either Dela- 
field’s haematoxylin or anilin blue. 

Land uses either aqueous, anilin, or alcoholic safranin, and 
uses the light green in clove oil, or in a mixture of clove oil and 
absolute alcohol. Make a saturated solution of light green in clove 
oil. Since the solution takes place slowly, the mixture should stand 
several days before using. If a small quantity of absolute alcohol 
be added to the clove oil, the stain dissolves more readily. For 
some structures the stain is more brilliant than with the simple 
clove-oil solution. 


90 


METHODS IN PLANT HISTOLOGY 


Sections from fresh material are fixed in 95 per cent alcohol; 
sections from preserved material are rinsed in alcohol or water before 
staining. The following schedule will summarize the method: 

1. Safranin, 2 to 24 hours. 

2. 50 per cent alcohol, until differentiated. 

3. Dehydrate in 95 and 100 per cent alcohol. 

4. Light green (in clove oil), 3 to 30 minutes. 

5. Xylol: 2 or 3 c.c. of absolute alcohol may be added to each 100 c.c. 
of xylol, if the free light green shows a tendency to precipitate. 

6. Mount in balsam. 

This stain is particularly good for phloem. Since the light green 
is not likely to overstain and does not extract the safranin, the 
combination is a rather easy one. Even the beginner can hardly 
fail to get a good preparation. 

Malachite Green and Congo Red.—I am indebted to Dr. Sharp 
for this method, which has been popular in Professor Gregoire’s 
laboratory at Louvain. 

Sections of fresh material should be treated with 95 per cent 
alcohol and then transferred to water. 

1. 3 per cent aqueous solution of malachite green or methylin blue, 
6 hours or more. 

2. Wash in water. 

3. Congo red, 1 per cent aqueous solution, 15 minutes. 

4. Wash in water. 

5. Rinse in 80 per cent alcohol. As soon as the malachite green or anilin 
blue color appears through the red, transfer quickly to 

6. Absolute alcohol. 

7. Xylol. 

8. Balsam. 

Iodine Green and Acid Fuchsin is another good combination 
for such sections. The stain will be particularly brilliant if sections 
from fresh material are fixed in 1 per cent chromo-acetic acid for 
10 to 24 hours; and then washed for an hour in water. Beginning 
with the sections in water, the procedure is as follows: 

Stain is aqueous iodine green for 12 to 24 hours. Then wash in 
water until the stain is nearly all washed out from the cellulose walls, 
but is still brilliant in the lignified walls. If the stain acts for too 
short a time, the washing-out process necessary to remove the stain 
from the cellulose walls will leave only a pale-green color in the ligni- 


FREEHAND SECTIONS 


91 


fied walls. Stain in aqueous acid fuchsin for 2 to 10 minutes. This 
should stain the cellulose walls sharply, but should not act long 
enough to affect the lignified tissues. Pour off the stain (which may 
be used repeatedly), and pour on 95 per cent alcohol, and imme¬ 
diately pour it off and add absolute alcohol. The 95 per cent alcohol 
should not act for more than 5 or 10 seconds, its only function being 
to save the more expensive absolute alcohol. From 10 to 30 seconds 
will usually be long enough for the absolute alcohol. Too long a 
period in the alcohols will weaken the stain. Clear in xylol or clove 
oil, and mount in balsam. 

If a 50 or 70 per cent alcoholic solution of iodine green has been 
used, the stain should be washed out in 50 per cent alcohol; otherwise 
the treatment is the same. 

Methyl Green (aqueous solution) and Acid Fuchsin is a good 
combination, and the student may find it easier to get a good differ¬ 
entiation than with iodine green. Follow the directions for the 
aqueous iodine green and acid fuchsin. It may be necessary to wash 
more rapidly, since the methyl green is easily extracted. 

Safranin and Gentian-Violet.—This is a good combination for 
vascular anatomy. Stain over night in safranin, rinse in 50 per cent 
alcohol until the stain is reduced to a light-pink color in the cellu¬ 
lose walls; then rinse in water and stain 5 to 10 minutes in aqueous 
gentian-violet. Rinse in water, dehydrate in 95 and 100 per cent 
alcohol, clear in clove oil, transfer to xylol and mount in balsam. 

Orange may be added to this combination, making a triple stain. 
In this case, do not reduce the safranin at all, but rinse quickly in 
50 per cent alcohol, then in water, stain in gentian-violet, rinse in 
95 per cent alcohol and stain for 1 or 2 minutes in orange dissolved 
in clove oil. This will not only reduce and differentiate the gentian- 
violet, but will reduce the safranin. Transfer to xylol and mount in 
balsam. If the safranin is drawn out too rapidly, stain for 15 to 30 
seconds in the orange, transfer to clove oil without any orange 
until the gentian-violet is satisfactory; then transfer to xylol and 
mount in balsam. 

The bordered pits of conifers, the Bars of Sanio, and the middle 
lamella are beautifully stained by this method. 

Other Combinations might be suggested, e.g., iodine green or 
methyl green with Bismarck brown, methyl green with Delafield’s 
haematoxylin; orange G might be added after the safranin and 


92 


METHODS IN PLANT HISTOLOGY 


Delafield’s haematoxylin, and various other stains might be tried. 
In double staining it is usually best to combine a basic with an acid 
stain. Green and red make a good contrast, but a section stained 
with iodine green and safranin would be a failure, because both 
stains would stain the xylem and neither would stain the cellulose. 
Both stains are basic. Red lignin and green cellulose could be secured 
by using safranin and acid green. Green lignin and red cellulose, 
as already indicated, can be got with iodine and acid fuchsin. 

The Time Required for the different processes varies greatly, 
and the time required for a subsequent process is often more or less 
dependent upon the time given to processes which preceded it. 
Good mounts of sections of the petiole of Nuphar advena have been 
secured from material which had been cut, fixed, stained in safranin 
and Delafield’s haematoxylin, and mounted in balsam, the entire 
time being less than 30 minutes. This is an extreme case, and 
nothing is gained, except time, and the saving of time is apparent 
rather than real, for the histologist always has something to do while 
the sections are in the stain. 

Preserved, Fresh, and Dry Material.—If sections are to be cut 
from material preserved in formalin, the piece should be washed in 
water, since the odor is annoying and the fumes are injurious to the 
eyes. 

The sections are placed in the stain from water. Sections from 
alcoholic material are transferred directly to the stain. If the 
material is in a mixture of alcohol and glycerin, the sections should 
be washed in water or 50 per cent alcohol until the glycerin has been 
removed before transferring to the stain. 

Some material cuts well when fresh, but cuts with difficulty when 
preserved. On the other hand, some material cuts well when pre¬ 
served, but hardly at all when fresh. Some material which is too soft 
to cut when fresh can be cut with ease after it has been in formalin 
alcohol for a week or more. 

Very hard material, like oak, hickory, maple, etc., should be 
boiled in water and treated with hydrofluoric acid before any section¬ 
ing is attempted. Cut the material into blocks suitable for sections 
and boil in water for several minutes; then transfer to cold water and, 
after several minutes, repeat the boiling. The alternate boiling and 
cooling, which should be repeated several times, drives out the air. 
Transfer to equal parts of commercial hydrofluoric acid and water. 


FREEHAND SECTIONS 


93 


From 1 to 3 weeks will be enough for most woods. Some oaks, ebony, 
apple, etc., may require a longer time and the acid may be used pure. 
One week in 25 per cent acid may be enough for corn stems. Wash 
thoroughly in water for a day or two. Then leave in equal parts of 
30 per cent alcohol and glycerin for several days before cutting. 
Material may be left indefinitely in the mixture of glycerin and 
alcohol. 

An article by Dr. La Dema M. Langdon, dealing with the prepara¬ 
tion and sectioning of hard, woody tissues, appeared in the Botanical 
Gazette of July, 1920. By her method, hard, woody tissues are 
softened so that they cut readily and can even be imbedded in paraffin 
successfully. 


OBJECTS MOUNTED WITHOUT SECTIONING 

Fern Prothallia, mounted without sectioning, make very useful 
preparations. Select desirable stages and fix in chromo-acetic acid 
for 10 to 24 hours; wash in water for 3 or 4 hours, changing the water 
frequently; stain in Delafield’s haematoxylin for 5 to 30 minutes; 
wash in slightly acidulated water for a few seconds, and then wash 
thoroughly in pure water. The prothallia must now be brought 
through a graded series of alcohols, 15,35,50,70,85,95, and 100 per cent 
being sufficiently close to prevent plasmolysis. Then use mixtures 
of alcohol and xylol, 3 parts absolute alcohol and 1 part xylol, 
2 parts alcohol and 2 parts xylol, 1 part alcohol and 3 parts xylol, 
and then pure xylol. Then bring the sections into a mixture of 
xylol and balsam, using at least 10 parts of xylol to 1 of balsam. If 
left in a shell, without corking, the xylol will soon evaporate, so that 
in a few days the prothallia may be mounted. Use the balsam in 
which the material has been standing, because any other balsam may 
have a different concentration. At every step in the process the 
prothallia should be examined under a microscope, so that any 
plasmolysis may be detected. If each succeeding step is tested with 
a single prothallium, a general disaster may be avoided. If plas¬ 
molysis takes place, weaken the reagent and try another prothallium. 
When a safe strength is found, bring on the bulk of the material, and 
use the same method with succeeding steps. The dangerous places 
are likely to be the transfer from alcohol to xylol and the transfer 
from xylol to balsam. The process is tedious, but the mounts are 
very firm and durable. The Venetian turpentine method is less 


94 


METHODS IN PLANT HISTOLOGY 


tedious and gives better results than have been secured by the 
method just described. Fix in the chromo-acetic-osmic mixture (d), 
described on page 26. Stain some prothallia in iron-alum haema- 
toxylin and some in Magdala red and anilin blue. When both have 
reached the thick turpentine, fine preparations can be made by 
mounting prothallia from both lots under the same cover. 

Sori of Ferns.—Instructive mounts of sori or of individual 
sporangia may be made without sectioning. It is better to choose 
ferns with thin leaves, since leaves thicker than those of Asplenium 
thelypteroides are likely to be unsatisfactory. If this fern is at hand, 
cut off several of the small lobes which bear three to six pairs of sori. 
Fix in chromo-acetic acid; wash in water; stain in Delafield’s 
haematoxylin, or omit staining altogether; pass through a series of 
alcohols, allowing each grade to act for at least 10 minutes; clear in 
clove oil, and mount in balsam. If the sori have begun to turn brown, 
better views of the annulus will be obtained without staining. 

Mosses and Liverworts.—Nearly all mounts are more successful 
by other methods, for which the student should consult the chapters 
on Bryophytes (chaps, xix and xx). Excellent mounts of the peristome 
of the moss can be made as follows: From fresh or preserved capsules 
cut off the peristome just below the annulus. Treat with 95 per cent 
alcohol 1 minute, absolute alcohol 2 to 5 minutes, clear in clove oil or 
xylol, and mount in balsam. It is a good plan to put at least three 
peristomes on a slide, one with the outside up, one with the inside up, 
and another dissected to show details of the teeth. 

Fairly good unstained mounts of the archegonia and antheridia 
of small mosses can be obtained by following the directions for 
mounting the sori of ferns. 

Beautiful and instructive mounts of the more delicate foliose 
Jungermanniaceae can be made by staining lightly in Delafield’s 
haematoxylin whole plants, or pieces as long as can be covered con¬ 
veniently. The method is that just given for fern prothallia. The 
mount should show both dorsal and ventral views. 

The Epidermis shows its best surface views without sectioning. 
Select some form with large stomata, like Lilium or Tulipa, strip 
pieces of epidermis from both sides of the leaf, and place them imme¬ 
diately in absolute alcohol for 1 or 2 minutes. Stain in Delafield’s 
haematoxylin; after this stain is satisfactory and all acid has been 
washed out, stain for 1 or 2 minutes in aqueous eosin, erythrosin, or 


FREEHAND SECTIONS 


95 


acid fuchsin; place directly into 95 per cent alcohol for a few seconds 
(merely to save the absolute alcohol), then into absolute alcohol for 
about 30 seconds, and then into clove oil. Mount in balsam. The 
epidermis is likely to curl and, unfortunately, patience seems to be 
the only remedy. In mounting, be careful to get pieces from both 
sides of the leaf, and be sure that the pieces are outside up. The 
inside of the epidermis is usually more or less rough, on account of 
the mesophyll torn off with it. Sedum purpurascens will show 
various stages in the development of stomata, even in epidermis 
stripped from mature leaves. The epidermis of the Sedums strips 
off very easily. If the large Sedum maximum is available, it is 
not difficult to strip off pieces 2 or 3 centimeters wide and several 
centimeters long. There is not much tendency to curl. The pieces 
may be spread out flat in a Petri dish, fixed in the chromo-acetic-osmic 
solution, just recommended for fern prothallia, or in this solution 
without the osmic acid. Wash in water, stain in Delafield’s haema- 
toxylin and eosin, or in safranin and gentian-violet. Then wash in 
water and run up through a series of alcohols—10, 20, 35, 50, 70, 85, 
and 95 per cent, about 2 minutes in each grade. If the gentian-violet 
is lost, stain for a minute in an alcoholic gentian-violet or in light 
green, for about 30 seconds; then transfer to clove oil, xylol, and 
balsam. This is a more tedious method, but it is worth the trouble. 
We have had the best results with the haematoxylin and eosin 
combination. 

Other Objects.—The cases just given will suggest other objects 
which might be mounted by such methods. Nearly all objects which 
used to be mounted in balsam without sectioning are now handled 
more successfully by the Venetian turpentine method. 


CHAPTER VII 
THE GLYCERIN METHOD 


It used to be an almost universal custom to mount unicellular 
and filamentous forms in glycerin or in glycerin jelly. The method 
is simple and easily mastered, but mounts must be sealed, and even 
when well sealed they do not long survive the ordinary use and abuse 
of the laboratory. However, some things are still mounted in glycerin 
or glycerin jelly. The greens and browns of moss protonema still 
keep much of their natural color in preparations which have been in 
use for many years. 

We have abandoned, completely, any mounting in glycerin; 
but the glycerin method, except the final mounting, precedes mounting 
in glycerin jelly, and it also constitutes the first part of the Venetian 
turpentine method. Consequently, it is very necessary to learn the 
capabilities and limitations of glycerin. 

The method, from fixing to mounting, as used in connection with 
staining and without staining, will now be described. 

Stained Preparations.—The familiar Spirogyra is a good form to 
begin with. Fix in the chromo-acetic-osmic solution described under 
(d) p. 26, or in this solution without the osmic acid, for 24 to 48 
hours. Wash in running water over night or 24 hours. If running 
water is not available, wash for 24 hours, changing the water 
frequently. 

The most satisfactory stain is Haidenhain’s iron-alum haematoxylin. 

Treat for 2 to 4 hours in a 2 per cent iron-alum; wash 10 to 20 
minutes in water, and then stain 4 hours, or over night, in J per cent 
haematoxylin. Wash in water 10 to 20 minutes and transfer to iron- 
alum again. This time the material must be watched under the 
microscope until the stain is just right. If the stain becomes weak 
in 2 or 3 minutes, the time in the stain was too short; if it re¬ 
quires an hour or more to reduce the stain to the proper tint, the 
time was too long. Different species of Spirogyra and even different 
collections, fixed in the same reagent, will differ in their reaction to 
stains; and different unicellular and filamentous forms, in different 

96 


THE GLYCERIN METHOD 


97 


fixing fluids, will present so much difference in times that only general 
suggestions can be given. When the stain is satisfactory, wash in 
running water for an hour. If this second iron-alum is not washed 
out thoroughly, its continued action will cause the preparation to fade. 

Put the material into 10 per cent glycerin (1 part glycerin and 
9 parts water), and then allow the water to evaporate gradually in a 
place as free from dust as possible. Nothing is better than a Petri 
dish for this purpose, because it presents a large surface for the evapo¬ 
ration of the water in the mixture. If there is much dust, cut a piece 
of filter paper just the size of the dish and let it float on the 10 per 
cent glycerin. The liquid will soak through the paper and evaporate 
without exposing the material itself to the dust. The process may 
be hastened, safely, by warming up to 35° C. The temperature of a 
paraffin bath—45° to 52° C.—causes such rapid evaporation that the 
material is likely to shrink. 

When the glycerin has become about as thick as pure glycerin, 
the material is ready for mounting. Place a small drop of glycerin, 
with the material, in the center of the slide, taking care not to put on 
so much that there will be a confusing tangle. Use scissors constantly 
so as not to injure filaments by trying to tease them out. Put on a 
round cover. There should be just enough glycerin to come to the 
edge of the cover-glass, but not any more , for it is impossible to seal 
a mount if glycerin has oozed out beyond the cover. 

The mount should now be sealed. Canada balsam, various 
asphalts, cements, flat varnish, gold size, and other things have been 
used. Canada balsam is always at hand and seems to be as good as 
any. Preparations which had been sealed with gold size more than 
fifty years before have been exhibited in perfect condition, but they 
must have been hidden away in some museum, for a glycerin mount 
would never survive fifty years of laboratory use. The gold size, as 
painters use it, is likely to be too thin for sealing mounts. Put some 
of it in a 1-ounce bottle with a wide neck and leave the cork out until 
the gold size thickens a little. Should it become too thick, thin it 
with turpentine. 

Nothing but practice will enable one to spin a good ring, but 
a good cameFs-hair brush, a good turntable, and a balsam neither 
too thick nor too thin will facilitate matters. Give the turntable a 
spin, and with the brush touch first the slide about as far out from the 
cover as you wish the ring to extend, then gradually approach the 


98 


METHODS IN PLANT HISTOLOGY 


cover. Dip the brush in the balsam again, and gradually extend 
the ring until it is about tV inch wide on the cover. The touch must 
be extremely gentle or the cover will be moved. Do not try to put 
on a thick ring the first time, but let a thin ring harden for an hour 
(months would do no damage), and then a thicker ring can be added 
without any danger. Thin rings are too likely to be broken, and 
thick rings are in the way if the preparation is to be examined with 
high powers. A medium ring is best, and it should consist of two 
coats, for a crack would seldom appear at the same place in both 
coats. A good shape and thickness for a ring are shown in Figure 20. 

I- ~~ - 1 


Fig. 20.—Slide, natural size, showing size and form of the ring 

The following is a summary of the foregoing processes: 

1. Fix in chromo-acetic-osmic acid, 24 to 48 hours. 

2. Wash in water, 24 hours. 

3. Iron solution, 2 hours. 

4. Wash in water, 10 minutes. 

5. ^ per cent haematoxylin, 3 to 24 hours. 

6. Wash in water, 10 minutes. 

7. Iron solution until stain is right. 

8. Wash in water, 1 hour. 

9. 10 per cent glycerin. 

10. Mount and seal. 

If the material has been fixed in formalin, it should be washed in 
water for 5 to 10 minutes before staining. Material preserved in 
70 per cent alcohol should be placed successively in 50 per cent, 35 per 
cent, 15 per cent alcohol, and then in water, allowing each to act 
for 15 to 30 minutes before being placed in the stain. 

Mayer’s haem-alum is also a good stain for filamentous algae and 
fungi which are to be mounted in glycerin. The process, after fixing 
and washing in water, is as follows: 

1. Transfer to the stain from water. 

It is seldom necessary to stain longer than 10 minutes. As a 
rule, it is better to dilute the stain (about 1 c.c. to 10 c.c. of distilled 
water) and allow it to act for 10 hours or over night. 

2. Wash in water, 20 minutes. 

3. 10 per cent glycerin until sufficiently concentrated. 

4. Mount and seal. 






THE GLYCERIN METHOD 


99 


Eosin is a good stain for many algae and fungi, when sharp out¬ 
lines rather than cell contents are to be brought out. After the 
material has been fixed and washed in water, stain in an aqueous 
solution of eosin for 12 to 24 hours. Pour off the eosin, which can be 
used repeatedly, and pour on a 1 or 2 per cent solution of acetic acid 
in water. Pour this off and pour on some more of the acid, until 
very little stain washes out. The process may require 2 to 5 minutes. 
Then place in 10 per cent glycerin containing about \ per cent acetic 
acid, and allow the glycerin to concentrate. The acetic acid is to 
prevent the stain from washing out. When the glycerin has reached 
the proper concentration, mount and seal as before. 

The following is a rapid method for forms like Eurotium and 
Penicillium: Fix in 100 per cent alcohol about 2 minutes; stain in 
aqueous eosin 5 minutes; wash in water about 1 minute; fix in 
1 per cent acetic acid 1 minute; then mount directly in 50 per cent 
glycerin to which about 1 per cent acetic acid has been added. It is 
hardly worth while to try this method with forms which have large 
cells; they are almost sure to collapse. If a form like Eurotium passes 
through the earlier processes without danger, but collapses when 
put into the 50 per cent glycerin, put it into the 10 per cent glycerin 
and allow the glycerin to concentrate. 

Mounting without Fixing or Staining.—Lt is sometimes desirable to 
retain the natural color of an obj ect. The chlorophyll green can usually 
be preserved by mounting directly in glycerin without any previous 
fixing. Other colors also are often preserved in this way. Moss 
protonema makes beautiful preparations by this method. If possible, 
select protonema showing the very young moss plants. The brown pro¬ 
tonema and brown bulbils preserve their color perfectly. Wash the dirt 
away from the protonema, which is then placed in 50 per cent glycerin. 
The brown or black spores of fungi are readily mounted in this way. 

The method is very useful when one finds a single specimen 
of Pediastrum , or any small form which would be lost in the more 
complicated processes. Place a large drop of 10 per cent glycerin 
on a slide; with a pipette, transfer the object to the drop, and allow 
the glycerin to concentrate. Then add a cover and seal the mount. 

GLYCERIN JELLY 

It is almost never necessary to mount anything in glycerin, because 
material can be transferred directly from glycerin to glycerin jelly. 


100 


METHODS IN PLANT HISTOLOGY 


If the glycerin jelly is well made, it is quite firm and mounts will last 
for a year or two, without sealing; but it is better to seal them with 
balsam. A very good formula is known as Kaiser’s gelatin. It is 
made as follows: One part by weight of the finest French gelatin 
is left for about 2 hours in 6 parts by weight of water; 7 parts of 
glycerin are added, and for every 100 grams of the mixture, 1 gram 
of concentrated carbolic acid. The whole is warmed for 15 minutes, 
stirring all the while until all the flakes produced by the carbolic 
acid have disappeared. Filter while warm through a fine-mesh 
cheese-cloth. 

To make a mount, take a small piece of the glycerin jelly, not 
more than half as large as a grain of wheat—the exact size will depend 
upon the material—warm it until it melts; and then transfer to it 
the material which has already been brought into thick glycerin. 
It is a good plan to touch the material to filter paper in order to 
remove as much glycerin as possible; for the less glycerin the firmer 
the mount will be. The mount may be sealed as soon as it is cool; 
but some prefer to let it stand for a week or two before sealing. In 
any case, it is a fairly firm mount, so that there is no danger of moving 
the cover. 

Everything which can be brought safely into pure glycerin can be 
mounted in glycerin jelly and the preparation is much more stable 
than a glycerin mount. 


CHAPTER VIII 

THE VENETIAN TURPENTINE METHOD 

Just as glycerin jelly is superior to glycerin as a mounting medium, 
so Venetian turpentine is superior to glycerin jelly. While the method 
was described by Pfeiffer and Wellheim 1 thirty years ago, it received 
no recognition in the United States or even in Europe. I made a 
casual trial of the method when preparing the first edition of this 
book, but the preparations were such miserable failures that the 
process did not seem worth mentioning. The method was next 
brought to my attention during a demonstration in Strasburger’s 
laboratory at Bonn. He was using preparations of Zygnema and 
Spirogyra, the staining of which surpassed anything I had ever seen. 
He remarked that it was not worth while to consult the lengthy 
article, because his preparations had been made by the authors and 
no one else had made a success of the method. However, when I 
returned, I made a careful study of the process, and finally learned 
to use it successfully. The details as given in this paper were too 
indefinite for practical use, but, after one has learned the method 
the article can be read with profit. 

The great practical advantages of the method are that prepara¬ 
tions are as hard and durable as balsam mounts, and that a much 
greater variety of staining is possible than in case of glycerin mounts. 

The principal features of the method are that material is brought 
from pure glycerin into alcohol, and from alcohol into Venetian 
turpentine, without passing through xylol or other clearing agent. 

After fixing and washing in water and staining in an aqueous 
stain, e.g., iron-alum haematoxylin, the process is as follows: 

1. 10 per cent glycerin until concentrated. 

2. Wash the glycerin out thoroughly in 95 per cent alcohol. 

3. Complete the dehydration in 100 per cent alcohol. 

4. 10 per cent Venetian turpentine in an exsiccator until the turpentine 
becomes thick enough for mounting. 

5. Mount in the Venetian turpentine. 

1 Ferdinand Pfeiffer and R.v. Wellheim, “Zur Preparation der Susswasseralgen,” Jahrbiichern 
fiir wissenschaftliche Botanik, 26 : 674-732, 1894. 

101 


102 


METHODS IN PLANT HISTOLOGY 


Even after the method became established, there occurred a 
period of several years during which it was practically impossible 
to get a Venetian turpentine suitable for histological use. Conse¬ 
quently, it was necessary to resort to glycerin jelly or to try various 
schemes for bringing material into Canada balsam; but good Venetian 
turpentines have appeared again and are even more satisfactory 
than those which established the method in popular favor. We have 
tested two brands which are giving uniform and excellent results: 
these are the “Venice Turpentine (True),” sold by the Fuller- 
Morrison Company, of Chicago; and the “Turpentine Venetian” 
(No. 2605), sold by the National Anilin and Chemical Company, of 
New York. There are probably other good turpentines and still 
others are likely to appear. Venetian turpentine is made from the 
resin of Larix europea. Since this resin is soluble in absolute alcohol, 
material can be transferred directly from absolute alcohol to a thin 
solution of the mounting medium, without the intervention of xylol 
or any other oil, thus eliminating the most dangerous stage in the 
preparation of filamentous algae and fungi and any forms which are 
to be mounted whole. 

The general outline, just given, is not sufficiently definite for a 
working introduction. The following concrete examples, describing 
the use of Venetian turpentine with an aqueous stain, with an alcoholic 
stain, and with a combination of aqueous and alcoholic stains, 
will be more practical than general directions. The steps from 
fixing to mounting, as used with an aqueous stain, will be described 
first, since this will introduce the method in its least complicated 
form. 

Haidenhain’s Iron-Haematoxylin.—-Using Spirogyra as a type, 
proceed as follows: 

1. Fix 24 hours in chromo-acetic acid. 


1 per cent chromic acid. 100 c.c. 

Glacial acetic acid. 3 c.c. 


The volume of the fixing agent should be at least 50 times that of 
the material to be fixed. 

2. Wash in running water 10 hours; if running water is not available, 
24 hours, changing often. 

3. 2 per cent aqueous solution of iron-alum (ammonia sulphate of iron), 
2 hours. 

4. Wash in running water, 20 minutes. If running water is not avail¬ 
able, wash in a large quantity of water and change frequently. 




THE VENETIAN TURPENTINE METHOD 


103 


5. Stain over night, or 24 hours, in \ per cent aqueous solution haema- 
toxylin. 

6. Wash in water, 20 minutes. 

7. 2 per cent aqueous solution of iron-alum, until the stain is satis¬ 
factory. This can be determined only by examining frequently 
under the microscope. 

8. Wash in water, 2 hours. If this washing is not thorough, the con¬ 
tinued action of the iron-alum will cause the preparations to fade. 

9. Transfer to 10 per cent glycerin, and allow the glycerin to concentrate 
until it has the consistency of pure glycerin. It is not necessary to 
use an exsiccator. Merely put the glycerin into shallow dishes, and 
leave it exposed to the air, but protected from dust. If the material 
is in Petri dishes or other dishes with a large surface, 3 or 4 days 
will be sufficient. This process may be hastened by warming, if 
the temperature does not go above 35° C. If the reduction from 
10 per cent glycerin to pure glycerin is accomplished in less than 
24 hours, the change in the concentration is so rapid that material is 
likely to suffer. 

10. Wash out the glycerin with 95 per cent alcohol. It will be necessary 
to change the alcohol several times. From 10 to 20 minutes will be 
sufficient if the alcohol is changed frequently. This alcohol cannot 
be used again for the same purpose, but it will be useful in cleaning 
one’s hands and in cleaning dishes which have contained Venetian 
turpentine. 

11. Complete the dehydration in 100 per cent alcohol: 10 minutes should 
be sufficient. 

12. Most failures are now ready to occur. 

From the absolute alcohol the material is transferred to a 
10 per cent solution of Venetian turpentine in absolute alcohol. The 
turpentine thickens as the alcohol evaporates, and when it reaches the 
consistency of pure glycerin the material is ready for mounting. 
The 10 per cent Venetian turpentine is very sensitive to moisture , and 
most failures are due to this characteristic; consequently the con¬ 
centration cannot be allowed to take place with the turpentine 
exposed to the air of the room. Use an exsiccator. This will not 
only absorb the moisture from the air, but will soon remove the 
alcohol from the turpentine mixture. Make an exsiccator as follows: 
Place a saucer full of soda lime (sodium hydroxide with lime) on a 
plate of glass, and cover with a bell jar. This is a simple and effective 
exsiccator. Instead, you may simply scatter soda lime in the bottom 


104 


METHODS IN PLANT HISTOLOGY 


of any low museum jar with tight-fitting cover. The tin cans, with 
tight covers, in which you get your pound of “ Improved Vacuum 
Coffee” make good exsiccators for small amounts of material. You 
may improvise other forms; the essential thing is to provide a small, 
air-tight place in which the soda lime may work. 

Instead of soda lime you may use fused calcium chloride or the 
white sticks of sodium hydroxide. 

We are now ready for the transfer from absolute alcohol to the 
10 per cent Venetian turpentine. Make the transfer quickly. Pour 
off the absolute alcohol and place the dish, with the material, in the 
exsiccator; then pour on the 10 per cent turpentine, and immediately 
put on the cover. This is better than to pour on the turpentine and 
then try to get the dish well placed in the exsiccator. 

The greater the surface of soda lime exposed, the more rapid 
will be the concentration of the Venetian turpentine. The con¬ 
centration must not be too rapid. Not less than 2 days should be 
allowed for the concentration of 30 c.c. of the turpentine in an 
ordinary Minot watch glass. 

Great care must be taken not to let any of the soda lime, or other 
drier, get into the turpentine. 

When the lime has become saturated, it may be heated until 
dry, and then used again. If material is put into an exsiccator with 
nearly saturated lime, the turpentine becomes milky. If the material 
is very valuable, wash in absolute alcohol until entirely free from 
any milky appearance, and start again in 10 per cent turpentine. 

As soon as the turpentine has attained the consistency of pure 
glycerin, it may be exposed to the air without any danger from 
moisture; but the turpentine would soon become too thick for 
mounting. If the turpentine has become too thick, thin it with 
a few drops of absolute alcohol or with 10 per cent or any thin solution 
of Venetian turpentine. 

Mount the material in a few drops of the Venetian turpentine 
and add a cover. Tapping on the cover with the handle of a needle 
or scalpel will often separate the filaments so that they are more 
convenient for examination. Square covers may be used since it is 
entirely unnecessary to seal the mounts, which are as hard and durable 
as those mounted in balsam. 

Material in the thickened Venetian turpentine, when not needed 
for immediate mounting, may be put into small bottles. The corks 


THE VENETIAN TURPENTINE METHOD 


105 


should be of the best quality; otherwise the turpentine will become 
too thick. While it can be thinned by adding thin turpentine, it is 
better, for easy mounting, not to let the turpentine become too thick. 
If the turpentine is only a little too thick, warming it gently will 
thin it enough for making mounts; but if any material is to be put 
away, a few drops of absolute alcohol or of a thin Venetian turpentine 
should be added. Material in Venetian turpentine, well corked 
and kept in the dark, does not fade or deteriorate in any way. 

Magdala Red and Anilin Blue.—Fix in chromo-acetic acid and 
wash in water, as described in the previous schedule. Transfer 
from water to 10 per cent glycerin and allow the glycerin to con¬ 
centrate. It is not necessary to use an exsiccator since there is no 
danger from moisture in the air. When the glycerin attains the 
consistency of pure glycerin, wash the glycerin out with 95 per cent 
alcohol and then proceed with the staining. 

1. Stain in Magdala red. At least two Magdala reds are sold 
by dealers. The one marked “echt” is more expensive, but, in our 
experience, is inferior to the one marked simply “Magdala red.” 
Make a 1 per cent solution in 90 per cent alcohol. We use the stain 
much stronger than recommended by Pfeiffer and Wellheim. The 
pure stain, allowed to act for 24 hours, does not stain too deeply. 

2. Rinse the material for a minute in 90 per cent alcohol. 

3. Stain in anilin blue, using a 1 per cent solution in 90 per cent 
alcohol. We prefer to make a fresh solution every time we have 
anything to stain. It is not necessary to measure it. A little of the 
powder—about half the bulk of a grain of wheat—in 30 c.c. of 90 
per cent alcohol, will give an efficient solution. The time required 
for successful staining will vary from 3 to 30 minutes. Do not put 
all the material into the anilin blue at once, but, by trying a few 
filaments at a time, find out what the probable periods may be. 

4. Rinse off the stain in 90 per cent alcohol, and then treat for a 
few seconds in acid alcohol (1 very small drop of HC1 to 30 c.c. of 90 
per cent alcohol). The acid alcohol fixes and brightens the anilin 
blue, but extracts the Magdala red. If the anilin blue or the acid 
alcohol acts for too short a time, the blue will be weak; if they act 
too long, the red is lost entirely. If the blue overstains too much, 
wash it out in 95 per cent alcohol. If the red overstains, wait until 
the mount is finished, and then reduce the red by exposing the slide 
to direct sunlight. 


106 


METHODS IN PLANT HISTOLOGY 


5. Absolute alcohol, 5 or 6 seconds. 

6. Transfer quickly to 10 per cent Venetian turpentine and pro¬ 
ceed as in the previous schedule. 

The surprising beauty of successful preparations will compen¬ 
sate for whatever failures may occur. Nuclei and pyrenoids should 
show a brilliant red, while the chromatophores and cytoplasm should 
be dark blue. The cell walls should show a faint bluish color. 

Haidenhain’s Iron-Alum Haematoxylin and Eosin.—Follow the 
schedule for iron-haematoxylin until the glycerin has been washed 
out in 95 per cent alcohol. Then stain for a minute in a solution of 
eosin in 95 per cent alcohol. Wash for a minute in 95 per cent alcohol, 
then a minute in absolute alcohol, and then transfer to the 10 per cent 
Venetian turpentine. 

Haidenhain’s Iron-Alum Haematoxylin and Safranin.—-Follow 

the schedule for iron-haematoxylin until the glycerin has been washed 
out in alcohol, and then add to the 95 per cent alcohol several drops 
of a solution of safranin in 95 or 100 per cent alcohol and allow the 
stain to act for 30 minutes or an hour. Then dehydrate in absolute 
alcohol and transfer to 10 per cent Venetian turpentine. 

Other Stains may be used. Aqueous stains should be used before 
starting with 10 per cent glycerin. Alcoholic stains should be in 
strong alcohol—about 90 per cent—and should be applied just after 
washing out the glycerin. 

This method is equally good for filamentous fungi and also for 
the prothallia of Equisetum and ferns, for delicate liverworts and 
mosses, and similar objects. 

If you have a good turpentine, good stains, and avoid moisture , 
the Venetian turpentine method should not be difficult, and the results 
with filamentous and unicellular forms and other small objects 
surpass anything yet secured by other processes. 


CHAPTER IX 


THE PARAFFIN METHOD 

The paraffin method is still the most important of all histological 
methods now in use. The results obtained by this method would 
have been regarded as almost miraculous by the histologists of one 
hundred years ago. At that time it was customary to observe 
things dry, and no cover-glasses were used. Section-cutting with 
sharp knives or razors did not become general until about 1830. The 
need for an instrument which would cut sections without demanding 
an extreme degree of manual dexterity was soon felt, but a successful 
microtome did not appear until much later. The latest microtomes, 
while rather complicated, give wonderful results. The Spencer 
microtome, shown in Figure 21, with the cooling attachment devised 
by Dr. Land, will cut even ribbons, 1 n in thickness, from such 
material as the antheridial receptacles of Marchantia. This means 
that a series of sections can be cut from pollen grains or spores too 
small to be seen by the naked eye. Many of the principles involved 
in this method are general in their application, and some of the pro¬ 
cesses are common to other methods. Before attempting the free¬ 
hand sectioning, the beginner should read the following paragraphs 
on killing and fixing, washing, hardening and dehydrating, and on 
clearing. 

KILLING AND FIXING 

As stated in the chapter on “Reagents” (chap, ii), the purpose 
of a killing agent is to bring the life-processes to a sudden termination, 
while a fixing agent is used to fix the cells and their contents in as 
nearly the living condition as possible. The fixing consists in so 
hardening the material that the various elements may retain their 
natural condition during all the processes which are to follow. Usu¬ 
ally the same reagent is used for both killing and fixing. Zoologists, 
from humane motives, may use chloroform for killing, while other 
reagents are used for fixing. In fixing root-tips, anthers, and other 
material for a study of mitotic figures, it is necessary that killing 
be very prompt. In a weak solution of chromo-acetic acid, nuclei 

107 


108 


METHODS IN PLANT HISTOLOGY 


which have begun to divide may complete the division, although the 
reagent might hinder nuclei from entering upon division. By 
treating for 20 minutes to 1 hour with Flemming’s weaker solution, 
or with a chromo-acetic solution containing a much smaller proportion 
of osmic acid, the killing will be greatly accelerated and the proportion 
of nuclei in division will be correspondingly greater. If filamentous 



Fig. 21. —Spencer rotary microtome with electric motor and Land’s apparatus for temperature 
control. 


algae are placed for 10 or 20 minutes in a chromo-acetic solution con¬ 
taining a little osmic acid, all the advantages of immediate killing 
will be secured. Material is then transferred to chromo-acetic acid 
containing no osmic acid. The short treatment with an osmic solu¬ 
tion is not likely to cause any serious blackening. 

Take the killing and fixing fluids into the field. If one waits 
until the material is brought to the laboratory there may be some 
fixing, but it will, in many cases, be too late to do much killing. 
Material which has begun to wilt is not worth fixing. Material like 





















THE PARAFFIN METHOD 


109 


Spirogyra, however, may be wrapped in several thicknesses of news¬ 
paper, placed in a botany can and brought into the laboratory. 
Before fixing, it should be placed in water for half an hour. Such 
forms suffer more from lack of air when placed in a bottle or a can 
than from lack of water when wrapped in wet newspaper. Branches 
with developing buds may be brought in and kept in water. Cones of 
the cycad, Ceratozamia , sent from Jalapa, Mexico, have arrived in 
Chicago with cell division still going on at a rapid rate. But such 
cases are extremes; as a rule, take the killing and fixing fluids into 
the field. 

Always have the material in very small pieces, in order that 
the reagents may act quickly on all parts of the specimens. Pieces 
larger than cubes of 1 cm. should be avoided whenever possible. 
While one sometimes needs sections 2 or even 3 cm. long, it is not 
likely to be necessary to fix pieces more than 4 or 5 mm. in thickness. 
For Very fine work no part of the specimens should require the 
reagent to penetrate more than 1 or 2 mm. 

For fixing agents of the chromic-acid series, the volume of the 
reagent should be about 50 times that of the material. 

Fixing agents with alcohol as an ingredient will fix a larger pro¬ 
portion of material. It must be remembered that the water, which 
is always present in living tissues, weakens the fixing agent. 

The time required for fixing varies with the reagent, the character 
of the tissue, and the size of the piece. About 24 hours is a com¬ 
monly recommended period for chromic-acid solutions, but 2 or even 
3 days will do no harm. 

Directions for making and using the various fixing agents are 
given in the chapters on “ Reagents” (chaps, ii, xxix). 

WASHING 

Nearly all fixing agents, except the alcohols, must be washed out 
from the material as completely as possible before any further steps 
are taken, because some reagents leave annoying precipitates which 
must be removed, and others interfere with subsequent processes. 
Aqueous fixing agents with chromic acid as their principal ingredient 
are washed out with water; aqueous solutions of corrosive sublimate 
are also washed out with water; but alcoholic solutions should be 
washed out with alcohol of about the same strength as the fixing 
agent; picric acid, or fixing agents with picric acid as an ingredient, 


110 


METHODS IN PLANT HISTOLOGY 


must not be washed out with water, but with alcohol, whether the 
picric acid be in aqueous or alcoholic solution. When washing with 
water, running water is best, and where this is not convenient the 
water should at least be changed frequently. The washing-out 
process usually requires about 24 hours. 

HARDENING AND DEHYDRATING 

After the material has been washed, it is necessary to continue 
the hardening and also to remove the water. Alcohol is used almost 
entirely for these purposes. It completes the hardening and at the 
same time dehydrates, that is, it replaces the water in the material, 
an extremely important consideration, for the least trace of moisture 
interferes seriously with the infiltration of the paraffin. 

The process of hardening and dehydrating must be gradual; 
if the material should be transferred directly from water to absolute 
alcohol, the hardening and dehydrating would be brought about 
in a very short time, but the violent osmosis would cause a ruinous 
contraction of the more delicate parts. In recent years, cytologists 
have been making the dehydration process more and more gradual. 
Twenty years ago most workers began the dehydration process with 
35 per cent alcohol and used the series 35, 50, 70, 85, 95, and 100 per 
cent alcohol. Some placed an intermediate grade between water and 
35 per cent alcohol. If plasmoylsis—the tearing away of the proto¬ 
plast from the cell wall—was avoided, the series was thought to be 
sufficiently gradual; but a series which may avoid plasmolysis may 
not be adequate if one is to study the finer details of cell structure. 
The following series is recommended: 2|, 5, 7§, 10, 15, 20, 30, 40, 50, 
70, 85, 95, and 100 per cent. There is no particular virtue in the 
fractions: it is convenient to make 10 per cent alcohol, dilute with 
an equal volume of water for the 5 per cent, and dilute the 5 per cent 
with an equal volume for the 2\ per cent. It will be noted that the 
series begins with very close grades and that the intervals are gradu¬ 
ally increased. The claim is that by beginning with very weak 
alcohols in close grades, more perfect dehydration can be secured 
at the end of the series. Various devices, like constant drip and 
osmotic apparatus, have been proposed to secure a more gradual 
transfer, but it is very doubtful whether these possess any real advan¬ 
tages. It is not necessary to use a large amount of alcohol: 2 or 3 
times the volume of the material is sufficient. 


THE PARAFFIN METHOD 


111 


The grades of alcohol may be used several times, but it must be 
remembered that pollen grains, fungus spores, starch grains, and vari¬ 
ous granules are likely to be left in the alcohol, so that it is wise to 
pour back through a filter each time, thus keeping the alcohols clean. 

As the alcohols absorb water from the material, they become 
weaker and weaker. If the various alcohols be poured in a large 
“waste alcohol” bottle, when a couple of liters has been accumulated, 
the strength may be determined by testing with an alcoholometer. 
Then any grade of less strength can be made from this stock. 

The time necessary for each of the stages has not been determined 
with any certainty. About 2 hours seems to be long enough for 
each of the grades from to 70 per cent; for 70, 85, and 95, about 
4 hours each; for absolute alcohol, 4 to 12 hours, changing 2 or 3 
times. If material is to be kept in alcohol, leave it in 85 per cent, 
but where labor is no object, it is better to go on and imbed 
it in paraffin. 

CLEARING 

Let us suppose that the material has been thoroughly dehydrated, 
so that not the slightest trace of water remains. If the supposition 
chances to be contrary to fact, all the work which has preceded, as 
well as all which is to follow, is only an idle waste of time. The 
purpose of a clearing agent is to make the tissues transparent, but 
clearing agents also replace the alcohol. At this stage the latter 
process is the essential one, the clearing which accompanies it being 
incidental. The clearing, however, is very convenient, since it 
shows that the alcohol has been replaced and that the material is 
ready for the next step. 

Various clearing agents are in use. Xylol is the most generally 
employed, and for most purposes it seems to be the best. Bergamot 
oil, cedar oil, clove oil, turpentine, and chloroform are used for the 
same purpose. Cedar oil and chloroform may, in some cases, be as 
good as xylol. 

Only a small quantity of the clearing agent is necessary, enough 
to cover the material being sufficient. 

The transfer from absolute alcohol to the clearing agent should 
be gradual, like the hardening and dehydrating processes. The most 
successful workers have been making this transfer more and more 
gradual. Thirty years ago it was customary to transfer from absolute 
alcohol directly to xylol; then a mixture of equal parts of absolute 


112 


METHODS IN PLANT HISTOLOGY 


alcohol and xylol was interpolated; in the second edition of this 
book three grades were placed between the absolute alcohol and 
xylol. It is undoubtedly better to make the transfer still more 
gradual. The following series seem to be safe, 2§, 5,10,15, 25, 50, 75, 
and 100 per cent xylol. These mixtures of absolute alcohol and xylol 
can be made with sufficient accuracy without measuring in a gradu¬ 
ate. The 50 per cent grade is made by mixing equal parts of abso¬ 
lute alcohol and xylol; the 25 per cent, by adding to the 50 per cent 
an equal volume of absolute alcohol; make the 10 per cent grade 
from the 25 per cent by adding a little more than an equal volume 
of absolute alcohol; in the same way, make the 5 per cent from the 
10 per cent, and the 2\ per cent from the 5 per cent. The different 
grades may be kept in bottles and may be used repeatedly. A couple 
of drops of safranin dissolved in absolute alcohol, added to the 50 
or 75 per cent xylol, will color the material a little and will often be 
helpful in orienting after the imbedding in paraffin. 

About 2 or 3 hours is enough for each grade. The pure xylol 
should be changed once or twice. Throughout the dehydrating and 
clearing it is a good plan to keep the material in Number 4 shells, 
which are made from glass tubing about 25 mm. in diameter. 

Other clearing agents may be used, but the process must be just 
as gradual. 

THE TRANSFER FROM CLEARING AGENT TO PARAFFIN 

This should also be a gradual process. The most convenient 
method is to place a small block of paraffin in the pure clearing agent 
with the material, but the block of paraffin should not rest directly 
upon the objects. Dr. Land uses coarse wire gauze, cut into strips 
about 15 mm. wide and tapered at both ends. The strip is then bent 
so that the pointed ends rest upon the bottom of the Number 4 shell, 
while the middle portion forms a flat table upon which the paraffin 
may rest. Dip the wire gauze table into xylol and then slip it care¬ 
fully into the Number 4 shell. The table portion should be 10 
or 15 mm. above the material, and there should be enough xylol 
to extend a few millimeters above the table. Place on the table a 
block of paraffin about equal to the volume of the xylol in the shell. 
The table not only prevents the paraffin from injuring the material by 
mechanical pressure, but insures considerable diffusion before the 
mixture of paraffin and xylol reaches the specimens. After 24 hours 


THE PARAFFIN METHOD 


113 


(or several days, if time permits) at room temperature, place the shell 
on a pasteboard box—slide boxes are good—on top of the paraffin 
bath. Do not place the shell directly upon the metal of the bath, 
since it is better to minimize heat. As soon as the paraffin is dis¬ 
solved, add some more, this time leaving the cork out, in order that 
the xylol may evaporate. About 24 hours on the top of the bath 
should be sufficient. 

THE PARAFFIN BATH 

This step is usually called infiltration, but when the transfer 
from the clearing fluid to paraffin is made gradually, as has just been 
indicated, the process of infiltration is already begun. It is now 
necessary to get rid of the xylol or other clearing agent. This is 
accomplished, to a considerable extent, by pouring off the mixture of 
xylol and paraffin and replacing it with pure melted paraffin. Pour 
off the pure paraffin immediately. This is important. You will 
notice that often, when the pure paraffin is poured on, a froth or 
scum will appear on the surface. Much of the xylol will be in this 
scum, and, if allowed to remain, it would diffuse into the mass and 
greatly prolong the time needed for infiltration. So, pour it off and 
add more pure paraffin, for some xylol remains in the tissues and must 
be removed. Do not put the shell into the bath, but use a flat dish of 
some sort. The main object is to have a fairly large surface exposed, 
so that the remaining xylol may evaporate as rapidly as possible. 
Change the paraffin 2 or 3 times. Soft paraffin (about 45° C.) may 
be used at first, but the second should be the paraffin of the grade in 
which the material is to be imbedded. If there are two baths, one 
should be kept at 46° C. and the other at 52° C., if the material is to 
be imbedded in 52° C. paraffin. While using the soft paraffin, keep the 
material in the 46° C. bath; for the harder paraffin, use the 52° C. bath. 
If there is only one bath, there is no object in using the 45° C. paraffin. 

Do not throw away the paraffin which you pour off, but put it in a 
waste jar or beaker, or, still better, in a small tin lard pail, in which 
you have made a lip to facilitate pouring. This can be placed in the 
bath, or, in winter, on the radiator, and the xylol will gradually 
evaporate. After long heating, the paraffin not only becomes as 
good as new, but even better, since it becomes more homogeneous 
and tenacious. If it contains dust or debris of any kind, it may be 
filtered with a hot filter. 


114 


METHODS IN PLANT HISTOLOGY 


The time required varies with the character of the material and 
the thoroughness of the dehydrating and clearing. If this schedule 
has been followed up to this point, the time will be much shorter than 
most investigators now deem necessary. Fern prothallia infiltrate 
perfectly in 15 to 20 minutes; onion root-tips in 20 to 30 minutes; 
ovaries of Lilium at the fertilization stage, 30 minutes to 1 hour; 
5 or 6 mm. cubes of endosperm of cycads, containing archegonia, 
2 to 2\ hours; median longitudinal sections, 4 or 5 mm. thick, through 
ovulate cones of Pinus Banksiana may require 6 or 8 hours; if serial 
sections through the entire cone are wanted, Miss Aase found that 
the time must be prolonged to 24 or even 48 hours. When one is 
dealing with many lots of the same kind of material, as in research 
work, the time required for infiltration is easily determined. As a 
rule, minimize heat. It is, probably, never necessary to use paraffin 
with a melting-point higher than 52° C. With Land’s cooling device 
sections 1 ju in thickness can be cut from 52° C. paraffin. 

IMBEDDING 

Material may be imbedded in paper trays, watch crystals, or any 
apparatus made for the purpose. The most satisfactory imbedding- 
dish we have used is a thin rectangular porcelain dish glazed inside. 
This dish, called a Verbrennungsschale, is made by the Konigliche 
Porzellan-Manufactur, Berlin, Germany. The most convenient 
sizes are 40X50X10 mm., 68X45X10 mm., and 91X58X15 mm. 
As listed, these dishes are not glazed; care should be taken to indicate 
that the dishes must be glazed inside (innen glasiert). Where these 
dishes are not available, any dishes of convenient size and shape can 
be used. The paper tray, if well made, is as good as anything. Thick 
ledger linen or thin, smooth cardboard make good trays. 

Smear the dish or tray with glycerin or soapy water to prevent 
sticking. Another way to prevent sticking is to put a piece of tissue 
paper in the dish, pour on water and make the tissue paper fit the inside 
of the dish, and then pour on the paraffin with the material to be im¬ 
bedded. The paraffin will not stick to the paper. If several objects are 
to be imbedded in one dish, it is best to have the dish as near the tem¬ 
perature of melted paraffin as possible; otherwise the objects may stick 
to the bottom, and it will be impossible to arrange them properly. 
Hot needles are good for arranging material. Great care should 
be taken not to have the dish too hot, since too high a temperature 


THE PARAFFIN METHOD 


115 


not only injures the material, but also prevents a thorough imbedding. 
Pour the paraffin with the objects into the imbedding-dish and arrange 
them so as to facilitate the future cutting-out from the paraffin cake. 
Look at Figures 22 and 23, representing the arrangement of root-tips 
in a paraffin cake. From a cake like that in Figure 22 it is easy to cut 
out tips for sectioning. The 
arrangement, or rather the 
lack of it, shown in Figure 23 
should be remembered only 
as an exasperating example. 

After the objects have 
been arranged, cool the cake 
rapidly by allowing the bot¬ 
tom of the dish to rest upon 
cold water. As soon as a 
sufficiently firm film forms 
on the surface of the cake, 
let water flow gently over 
the top. After the cake has 
been under water for a few 
minutes, the paraffin will 
either come out and float 
on the water or, at least, it 
will be easily removed from 
the dish. If paraffin cools 
slowly it crystallizes and 
does not cut well. The layer 
of paraffin should be just 
thick enough to cover the 
objects, not only as a matter 
of economy, but because a thick layer retards the cooling. Very 
small objects, like the megaspores of Marsilea, ovules of Silphium, 
etc., may simply be poured out upon a cool piece of glass, which has 
been smeared with glycerin or soapy water. In this way, thin cakes 
are made which harden very rapidly. 

If one is doing much imbedding, it is worth while to have the par¬ 
affin cakes uniform in size and to have a convenient method of filing. 
In our own collection, there are more than 6,000 paraffin cakes. They 
are filed in pasteboard boxes 28 cm. long, 10 cm. wide, and 2 cm. deep. 


imrmw 
mm vvvvrir 
frmimrr 

Fig. 22 



Fig. 23 

Figs. 22, 23. —Paraffin cakes of root-tips, the upper 
(Fig. 22) showing a good arrangement, the lower (Fig. 23) 
showing fewer tips and most of these not in position to 
be blocked without injury. 







116 


METHODS IN PLANT HISTOLOGY 


With the generic name written on the box, it is easy to find anything 
in the large collection. 

CUTTING 

As soon as the paraffin is thoroughly cooled, it is ready for cutting. 
Trim the paraffin containing the object into a convenient shape, and 
fasten it upon a block of wood. Blocks of pine f inch long and f inch 
square are good for general purposes. Put paraffin on the end of the 
block so as to form a firm cap about J inch thick. Warm the cap 
and the bottom of the piece containing the object, and press them 
lightly together; then touch the joint with a hot needle, put the 
whole thing into cold water for a minute, and it is ready for cutting. 
Cutting can be learned only by experience, but a few hints may not 
come amiss: 

a) Keep the knife sharp. There should be two hones, one for 
use when the knife is- rather dull and the other for finishing. For 
the first hone, nothing equals a fine carborundum hone. About 
5.5X22.5 cm. is a good size. A hard Belgian hone, of the same 
size, may be a little better for finishing. Flood the stone with water, 
and rub it with the small slip which accompanies all high-grade hones; 
this not only makes a lather which facilitates the sharpening, but 
it also keeps the surface of the hone flat. As soon as the edge of the 
knife appears smooth and even under a magnification of 30 or 40 
diameters, the sharpening is completed with a good strop. It is 
better to sharpen the knife every time you use it. A first-class 
microtome knife, in perfect condition, is unsurpassed for cutting 
paraffin sections, but it requires both time and skill to keep the edge 
perfect. More than 20 years ago we began to experiment with the 
Gillette safety-razor blade and devised several holders for it, some 
of them more or less successful. Mr. Strickler finally perfected a 
holder which has already been mentioned. In using this holder 
the blade should not project more than 1 mm. The Gillette blade is 
harder than the ordinary microtome knife, and is so sharp that the 
edge appears smooth, even under a high-power dry lens. The bevel 
is about the same as that of a microtome knife which has been 
“backed” for sharpening. With this blade in the holder as made 
by Mr. Strickler or Mr. Larsen, we have cut smooth ribbons of 
Selaginella strobili, sections through the sporangium region of the 
whole plant of Isoetes, sections of stems of Cucurbita, in fact, we have 
not used an ordinary microtome knife for cutting paraffin ribbons for 


THE PARAFFIN METHOD 


117 


more than 15 years. Many fail at the first attempt and go back to 
the continual drudgery of sharpening microtome knives. If the 
holder is placed in the microtome at the angle used for a microtome 
knife, failure is certain, because the blade will scrape rather than cut. 
The angle should be considerably nearer vertical than in the case of a 
microtome knife. A study of Figure 6 should enable anyone to secure 
the proper angle. 

b) Keep the microtome well oiled and clean. 

c) Trim the block so that each section shall be a perfect rectangle. 



Fig. 24.—The ribbon 


CC 


A ribbon of sections like that shown in Figure 24 A is much better 
than one like B of the same figure, because sections will usually come 
off in neater ribbons if the knife strikes the longer edge of the rectangle, 
so that the sections are united by the longer sides rather than by the 
shorter. Crooked ribbons are caused by wedge-shaped sections, and 
are always to be avoided, because they 
make it difficult to economize space, 
and also because they present such 
a disorderly appearance. The knife, 
which should be placed at a right 
angle to the block and not obliquely, 
should strike the whole edge of the 
block at once, and should leave in the 
same manner. 

If sections stick to the knife, it 
may be that the knife is too nearly 
parallel with the surface of the block, as in Figure 25 A. By inclining 
the knife as in Figure 25 B, this difficulty is often obviated. A split or 
scratch in the paraffin ribbon may be caused by a nick in the knife. 
Use some more favorable position of the edge, or sharpen the whole 



A B 

Fig. 25.—Position of knife 





















118 


METHODS IN PLANT HISTOLOGY 


knife. A split or a scratch in the ribbon is often caused by some 
hard granule which becomes fastened to the inner side of the edge of 
the knife. This is the most common cause of the difficulty. Simply 
wipe the knife by an upward stroke of the finger, slightly moistened 
with xylol. Do not use a cloth. 

Sometimes good sections can be cut with a rather slow stroke 
when a rapid stroke fails. When paraffin is rather hard, sections 
may sometimes cut nicely at 5 when, at 10 n , ribbons cannot be 
secured. If very thin sections are desired and the paraffin seems 
too soft, cool the paraffin and the edge of the knife with ice, by Land’s 
cooling device, or by pressing a piece of ice against the paraffin block 
and the knife. Sometimes hard paraffin does not ribbon well. This 
difficulty may be remedied by dipping a hot needle in soft paraffin 
and applying it to the opposite edges of the block to be cut. Often 
the mere warming of the opposite edges of the block with a hot needle 
is sufficient. 

Another method, suggested by Dr. Land to facilitate the cutting 
of difficult material, has been tested in this laboratory and has been 
found to be very effective. Paraffin absorbs a small amount of water, 
or water penetrates between the crystals of paraffin. At any rate, 
water reaches cell walls and, perhaps, other structures which have 
not been completely infiltrated, and thus softens them. The paraffin 
cakes may be left for weeks in water. Cakes of class material may 
be put in water in a fruit can and kept until ready for use. After 
such treatment, smooth ribbons may be cut from material which 
would hardly cut at all without it. 

A ribbon carrier is very convenient. A good carrier can be made 
by mounting a couple of spools 15 or 20 inches apart, with a strong 
piece of cloth for a band. More elaborate carriers may be made if 
one has tools. 


FIXING SECTIONS TO THE SLIDE 

Mayer’s Fixative.—Sections must be firmly fixed to the slide, 
or they will be washed off during the processes involved in staining. 
Mayer’s albumen fixative is excellent for this purpose. Formula: 


White of egg (active principle). 50 c.c. 

Glycerin (to keep it from drying up). 50 c.c. 

Salicylate of soda (antiseptic, to keep out bac¬ 
teria, etc.) . 1 g. 





THE PARAFFIN METHOD 


119 


Shake well and filter through cheese-cloth. It will keep from 2 to 
6 months, but, to say the least, it is never better than when first made 
up. Of course, white of egg may be used alone, since the other two 
ingredients are merely incidental. Put a small drop of fixative on 
the slide, smear it evenly over the surface, and then wipe it off with 
a clean finger until only a scarcely perceptible film remains; then 
add several drops of distilled water and float the sections or ribbons 
on the water. Warm gently until the paraffin becomes smooth and 
free from wrinkles. Wrinkled or curved ribbons may be straightened 
by touching with a needle at each end and pulling gently, just as the 
ribbon begins to smooth out in the warming. Be careful not to melt 
the paraffin, for the albumen of the fixative coagulates with less 
heat than is required to melt the paraffin. If the paraffin should 
melt, run some cold water under it, and transfer the ribbon to another 
slide, prepared with fixative and water. It is a very good plan to 
put the slide on a metal bath or warming plate, like that shown in 
Figure 11. After the sections have become smooth, remove the 
surplus water and leave them on the bath with a couple of thicknesses 
of blotting paper under them for 3 or 4 hours, or, better, over night. 
If the fixative is used alone, as is often the case when sections are 
very thick, none of the delay is necessary, since the sections are 
merely laid upon the fixative and pressed down gently with the 
finger. 

Land’s Fixative.—Mayer’s fixative is so easily prepared and it 
keeps so well that it is in universal use; but, in many cases, it will 
not hold the section to the slide. Moss archegonia and moss capsules 
are likely to wash off, especially if cut rather thick. Large sections 
of cones of conifers are almost sure to float off as soon as the slide 
comes into the xylol or alcohol. Sections of ovules of cycads, as 
soon as they attain a length of 1.5 to 2 cm., are likely to wash off. 
For handling these more difficult cases, Dr. Land devised a fixative 
which has proved satisfactory, even in such extreme cases as sections 
of ovulate cones of Pinus Banksiana 2 cm. long. Formula: 


Gum arabic. 1.0 g. 

Bichromate of potash. 0.2 g. 

Water. 100.0 c.c. 


The mixture will not keep; the formula is given merely to indicate 
its composition. Make a 1 per cent solution of gum arabic in water, 





120 


METHODS IN PLANT HISTOLOGY 


which will keep as well as Mayer’s fixative; but make the bichromate 
solution immediately before using. Do not make the solution 
stronger than 1 per cent; usually 0.2 per cent is strong enough. 
Dr. Land does not measure, but simply adds enough bichromate 
crystals to make the water pale yellow. 

Smear a few drops of the 1 per cent solution of gum arabic on the 
slide; flood with the bichromate solution; warm to straighten the 
ribbons; drain off the excess water and let the preparation dry in 
the light. The exposure to light renders the gum insoluble in water. 
LePage’s glue or Mayer’s albumen fixative may be used instead of 
gum arabic. 

The foregoing directions are taken from Dr. Land’s notes. 

With the ordinary Mayer’s albumen fixative the bichromate of 
potash, without the gum arabic, may be used in floating out ribbons, 
and makes a stronger fixative than the Mayer’s formula. 


Szombathy’s Fixative.— 

Gelatin... 1 g. 

Distilled water. 100 c.c. 

Salicylate of soda (a 2 per cent solution). 1 c.c. 

Pure glycerin. 15c.c. 


Dissolve the gelatin in water at 30° C., add the salicylate of soda, 
shake well, cool, and filter through cheese-cloth; then add the 15 c.c. 
of glycerin. The solution should be perfectly clear. 

A couple of drops of the fixative,.with a couple of drops of 2 per cent 
formalin, is rubbed on the slide. The sections are then added, and 
straightened out. The formalin makes the gelatin insoluble. The 
fixative is much like Land’s and is used for difficult material which is 
not held by Mayer’s fixative. 

REMOVAL OF THE PARAFFIN 

To remove the paraffin, place the slide in a Stender dish of xylol. 
About 5 minutes will be sufficient for sections 10 u thick. The time 
may be shortened a little by gently warming the slide. Never heat 
the slide enough to melt the paraffin. Never attempt to warm the paraf¬ 
fin over a lamp. Overheating is ruinous. 

Many prefer to remove the paraffin by pouring on xylol or 
turpentine. Hold the slide at an angle of 45°, and pour on a little 
xylol or turpentine. If the slide has been slightly warmed this 
should carry off the paraffin immediately. The reagent used in 






THE PARAFFIN METHOD 


121 


this first pouring cannot be used again. Now flood the slide several 
times with the turpentine or xylol, pouring the reagent back into 
the bottle. 

REMOVAL OF XYLOL OR TURPENTINE 

To remove the xylol, place the slide in equal parts of xylol and 
absolute alcohol in a Stender dish. After 5 minutes, transfer to 
absolute alcohol, which should also be allowed to act for 5 minutes. 

If the pouring process is preferred and turpentine has been used 
to remove the paraffin, remove the turpentine by flooding the slide 
with 95 per cent alcohol. About 100 c.c. of turpentine and 200 c.c. 
of 95 per cent alcohol should be sufficient for 50 slides, even if the 
sections are to be mounted under the longest covers. By keeping 
both reagents in bottles and pouring the liquid on the slide, the 
reagents are always fresh. A given quantity of the reagent will 
prepare as many slides by one method as by the other. 

TRANSFER TO THE STAIN 

After the paraffin has been removed with xylol or turpentine, 
and the xylol or turpentine has been rinsed off with alcohol, the next 
step is the staining. If the stain is a strong alcoholic one (85 to 100 
per cent alcohol), transfer directly to the stain. If the stain is in 
70 per cent alcohol, pass through 95 and 85 per cent alcohol, 2 minutes 
in each, before staining. If an aqueous stain is to be used, pass down 
the whole series—95, 85, 70, 50, 35, and water—2 minutes in each, 
before placing the slide in the stain. 

This is rather tedious, but, for cytological work, it seems to be 
necessary. For general morphological work, the slide may be 
transferred directly from the absolute or 95 per cent alcohol to any 
stain. 

DEHYDRATING 

After the sections have been stained, they must be dehydrated. 
If they have been stained in a strong alcoholic solution, transfer to 
95 and then to 100 per cent alcohol, 2 minutes in each, if the stain 
does not wash out too rapidly. If stained in an aqueous solution, 
pass-through the series, water, 35, 50, 70, 85, 95, and 100 per cent 
alcohol, about 2 minutes in each. 

With stains which wash out rapidly, the times must be shortened 
and some of the alcohols must be omitted. With aqueous gentian- 
violet, all must be omitted except the 95 and 100 per cent, and even 
in these the time must be shortened to a few seconds. 


122 


METHODS IN PLANT HISTOLOGY 


CLEARING 

After the sections have been dehydrated, they must be cleared, 
or made transparent by some clearing agent. The clearing agent 
must be a solvent of balsam, but it is not at all necessary that the 
balsam shall be dissolved in the particular clearing agent which has 
been used. Xylol balsam is used not only when preparations have 
been cleared in xylol, but also when they have been cleared in clove 
oil, cedar oil, bergamot oil, or other clearing agents. 

Xylol is the most generally useful clearing agent. Place the 
slide in equal parts of xylol and absolute alcohol and then in pure 
xylol, allowing each to act for about 2 minutes. 

Clove oil is also an excellent clearing agent. The clove oil should 
follow the absolute alcohol, without any mixtures. Pour on a few 
drops of clove oil, and drain them off at once in such a way as to carry 
with them whatever alcohol may still remain. Then flood the slide 
repeatedly with clove oil, draining the clove oil back into the bottle. 
If judiciously used, 50 c.c. of clove oil is enough to clear 100 prepara¬ 
tions. Sections are usually cleared in a few seconds. The only 
objection to clove oil is that mounts harden slowly. To overcome 
this difficulty, the slide should be dipped in xylol for a minute before 
mounting in balsam. 

Synthetic oil of wintergreen is much less expensive and some 
claim that it is just as good as clove oil. 

For clearing sections on the slide, other clearing agents are 
hardly worth mentioning. 

MOUNTING IN BALSAM 

After the sections are cleared, wipe the slide on the side which 
does not bear the sections. Put on a drop of Canada balsam and 
add a clean, 1 thin cover. Before the cover is put on, pass it through 
the flame of an alcohol lamp to remove moisture, for it would be a pity 
indeed to injure a preparation at this stage of the process. Add a 
label, and the mount is complete. 

1 Slides and covers should be treated with hydrochloric acid, or equal parts of hydrochloric 
acid and water, for several hours. They should then be thoroughly rinsed in water and' wiped 
with a cloth perfectly free from lint. After rinsing in water, they may be kept in 95 per cent 
alcohol. When a cover is needed for use, it is Dr. Land’s practice to rest the corner of the cover 
on a piece of filter paper to remove the drop of alcohol; then pass the cover through the flame of a 
Bunsen or alcohol lamp. The film of alcohol will burn and the cover may warp, but it will usually 
straighten, and it will be clean and dry. 

The mixture of sulphuric acid and bichromate of potash, used for cleaning laboratory glass¬ 
ware, is equally good for slides and covers. 


THE PARAFFIN METHOD 


123 


A TENTATIVE SCHEDULE FOR PARAFFIN SECTIONS 
It will be useful to give several tentative schedules for the use 
of beginners. It cannot be too strenuously insisted that these schedules 
are only tentative, their sole object being to give the beginner a start. 
The following is a tentative schedule for the ovary of a lily at any 
period before fertilization. The pieces should not be more than 
12 mm. in length. 

1. Chromo-acetic acid, 1 day. 

2. Wash in water, 1 day. 

3. 2\, 5, 10, 15, 20, 30, and 50 per cent alcohol, 3 hours each; 70, 85, 
and 95 per cent alcohol, 4 hours each; absolute alcohol, 6 to 10 hours, 
changing 2 or 3 times. 

4. Mixtures of absolute alcohol and xylol; 2\, 5, 10, 15, 25, 50, 75, and 

100 per cent xylol, 2 or 3 hours in each grade. Change the pure 
xylol once or twice. N 

5. Paraffin and xylol, 48 hours. 

6. Melted paraffin in the bath, 30 to 40 minutes. 

7. Imbed. 

8. Section; about 10 y is a good thickness. 

9. Fasten to the slide. 

10. Dissolve off the paraffin in xylol, 5 minutes. 

11. Xylol and absolute alcohol, equal parts, 2 minutes; 100, 95, 85, and 
70 per cent alcohol, 2 minutes each. 

12. Stain in safranin (alcoholic), 6 hours or over night. 

13. Rinse in 50 per cent alcohol, using a trace of HC1 if necessary; then 
in 70, 85, 95, and 100 per cent alcohol, 2 minutes each. 

14. Stain in gentian-violet dissolved in clove oil (or in clove oil with a 
little absolute alcohol), 10 minutes. 

15. Treat with pure clove oil until the gentian-violet stain is satisfactory. 

16. Rinse in xylol, 1 minute. 

17. Mount in balsam. 

18. Label. 

That the paraffin method is tedious and complicated is univer¬ 
sally recognized. Many substitutes have been tried, but without 
enough success to justify even a reference. 


CHAPTER X 

THE CELLOIDIN METHOD 

The celloidin method has almost disappeared from botanical 
microtechnic, because material too hard for imbedding in paraffin 
can be cut without any imbedding at all, and material too delicate 
to be cut without a supporting medium can be imbedded in paraffin. 
But these two categories do not cover all the ground; celloidin still 
has its advantages. Years ago, a piece of rotten wood from an ancient 
Egyptian mummy case was brought to the writer for identification. 
It could be rubbed into powder in the fingers, and had to be handled 
gently to keep it from falling to pieces. It was cut very successfully 
in celloidin. Stems too hard for paraffin may be cut in celloidin 
when it is desired to preserve cell contents. Celloidin is still very 
valuable for most of the sections used in medical schools, because the 
sections can be prepared in great numbers and each student can take 
a section, add a drop of balsam and a cover, and have a preparation 
of his own ready to study. Where serial sections are necessary, 
as in most morphological and cytological work, the method is too 
tedious to be worth even a trial, unless the sections cannot be cut 
in any other way. Besides, most of the more valuable stains color 
the celloidin matrix, and if the matrix be removed, the more delicate 
elements may be displaced or even lost. 

Celloidin and collodion are forms of nitro-cellulose. They are 
inflammable, but do not explode. Schering’s celloidin, which is only 
a collodion prepared by a patented process, is in general use for 
imbedding. Granulated and shredded forms of celloidin are on the 
market, but the tablets are more convenient. Directions for making 
the various solutions accompany the celloidin. To make a 2 per cent 
solution, add to 1 tablet enough ether-alcohol to make the whole 
weigh 2,000 g. To make a 4 per cent solution, add another tablet, 
and to make a 6 per cent solution, add an additional tablet, and so on. 

The collodion method was published by Duval 1 in 1879. Cel¬ 
loidin was recommended by Merkel and Schiefferdecker 2 in 1882. 

1 Duval, Journal de Vanatomie, 1879, p. 185. 

2 Merkel and Schiefferdecker, Archiv fiir Anatomie und Physiologie, 1882. 

124 


THE CELLOIDIN METHOD 


125 


The principal features of the method are as follows: Material is 
dehydrated in absolute alcohol; treated with ether-alcohol; infil¬ 
trated with celloidin; imbedded in celloidin; hardened in chloro¬ 
form or alcohol; after which it is cut, stained, and mounted. 

Eycleshymer, who brought the celloidin method to a high degree 
of efficiency, published in 1892 a short account, which may be sum¬ 
marized as follows: Put the celloidin tablet, or fragments, into a 
wide-mouthed bottle, and pour on enough ether-alcohol (equal parts 
ether and absolute alcohol) to cover the celloidin. Occasionally 
shake and add a little more ether-alcohol until the celloidin is all 
dissolved. The process may require several days. The solution 
should have the consistency of a very thick oil. Label this solu¬ 
tion Number 4. Solution Number 3 is made by mixing 2 parts of 
solution Number 4 with 1 part of ether-alcohol. Solution Number 2 
is made by mixing 2 parts of Number 3 with 1 part of ether-alcohol. 
Solution Number 1 consists of equal parts of ether and absolute 
alcohol. 

After dehydrating, the material is placed successively in solutions 
1, 2, 3, and 4. For an object 2 mm. square, 24 hours in each solu¬ 
tion is sufficient; for the brain of a cat, a week is not too long. 

A paper tray may be used for imbedding. Pour the object, with 
the thick solution, into the tray and harden in chloroform for 24 hours; 
then cut away the paper and place the block in 70 per cent alcohol 
for a few hours. The material may be left indefinitely in a mixture 
of equal parts of 95 per cent alcohol and glycerin. 

Before cutting, the object is mounted upon a block of wood. A 
block, suited to the microtome clamp, is dipped in ether-alcohol, 
which removes the air and insures a firmer mounting. Dip the end 
of the block of wood in solution Number 3, and the piece of celloidin 
containing the object in solution Number 1. Press the two firmly 
together, and place in chloroform until the joint becomes hardened. 

Set the blade of the microtome knife as obliquely as possible. 
Both the object and the knife should be kept flooded with 70 per cent 
alcohol, and the sections, as they are cut, should be transferred to 
70 per cent alcohol. 

Stain in Delafield’s haematoxylin for 5 to 30 minutes. Wash in 
water for about 5 minutes, and then decolorize in acid alcohol (2 to 5 
drops of hydrochloric acid to 100 c.c. of 70 per cent alcohol) until 
the stain is extracted from the celloidin, or at least until the celloidin 


126 


METHODS IN PLANT HISTOLOGY 


retains only a faint pinkish color. Wash in 70 per cent alcohol (not 
acid) until the characteristic purple color of the haematoxylin replaces 
the red due to the acid. Stain in eosin (preferably a 1 per cent solu¬ 
tion in 70 per cent alcohol) for 2 to 5 minutes. Dehydrate in 95 per 
cent alcohol for about 5 minutes. Absolute alcohol must not be 
used, unless it is desirable to remove the celloidin matrix. Eycleshy- 
mer’s clearing fluid (equal parts of cedar oil, bergamot oil, and 
carbolic acid) clears readily from 95 per cent alcohol. Mount in 
balsam. 

If serial sections are necessary, arrange the sections upon a slide, 
using enough 70 per cent alcohol to keep the sections moist, but not 
enough to allow them to float. Cover the sections with a strip of 
toilet paper, which can be kept in place by winding with fine thread. 
After the sections have been stained and cleared, remove the excess 
of clearing fluid by pressing rather firmly with a piece of blotting- 
paper. Then remove the toilet paper and mount in balsam. 

With occasional slight modifications, we have used the method 
as presented by Eycleshymer in his classes. Instead of the graded 
series of celloidin solutions, we use a 2 per cent solution, which is 
allowed to concentrate slowly by removing the cork occasionally, 
or by using a cork which does not fit very tightly. The material 
is imbedded when the solution reaches the consistency of a very 
thick oil. If the material is to be cut immediately, we prefer to 
imbed it and fasten it to the block at the same time. The blocks 
should have surface enough to accommodate the objects, and should 
be about J inch thick. White pine makes good blocks; cork is 
much inferior. Place the block for a moment in ether-alcohol and 
then dip into the 2 per cent celloidin the end of the block which was 
left rough by the saw. With the forceps remove a piece of the 
material from the thick celloidin and place it upon the block, taking 
care to keep it right side up. Dip the block with its object first in 
thick celloidin, then in thin, and after exposing to the air for a few 
minutes drop it into chloroform, where it should remain for about 
10 to 20 hours. It should then be placed in equal parts of glycerin 
and 95 per cent alcohol, where it may be kept indefinitely. If the 
material is hard, like many woody stems, it will cut better after 
remaining in this mixture for a couple of weeks. 

The following schedules, beginning with the celloidin sections 
in 70 per cent alcohol, will give the student a start in the staining: 


THE CELLOIDIN METHOD 


127 


Delafield's Haematoxylin and Eosin.—• 

1. 70 per cent alcohol, 2 to 5 minutes. 

2. Delafield’s haematoxylin, 5 to 30 minutes. 

3. Wash in water, 5 minutes. 

4. Acid alcohol (1 c.c. hydrochloric acid+100 c.c. of 70 per cent alcohol) 
until the stain is extracted from the celloidin, or at least until only a 
faint pinkish color remains. 

5. Wash in 70 per cent alcohol (not acid) until the purple color returns. 

6. Stain in eosin (preferably a 1 per cent solution in 70 per cent alcohol), 
2 to 5 minutes. 

7. Dehydrate in 95 per cent alcohol, 2 to 5 minutes. Do not use abso¬ 
lute alcohol unless you wish to dissolve the celloidin, which is not 
necessary with this staining. 

8. Clear in Eycleshymer’s clearing fluid, usually 1 to 2 minutes, but 
sometimes 5 to 10 minutes. 

9. Mount in balsam. 

Safranin and Delafield's Haematoxylin.— 

1. 70 per cent alcohol, 2 to 5 minutes. 

2. Safranin (alcoholic), 6 to 24 hours. 

3. Acid alcohol (a few drops of hydrochloric acid in 70 per cent alcohol) 
until the safranin is removed from the cellulose walls. 

4. Wash in 50 per cent alcohol, 5 to 10 minutes to remove the acid. 

5. Delafield’s haematoxylin, 2 to 5 minutes. 

6. Wash in water, 5 minutes. 

7. Acid alcohol, a few seconds. 

8. Dehydrate in 95 per cent alcohol, 2 to 5 minutes, then in absolute 
« alcohol, 2 to 5 minutes, which will partially dissolve the celloidin. 

9. Clear in clove oil, which will complete the removal of the celloidin. 

10. Be sure that the sections are free from fragments of celloidin and then 

mount in balsam. 

Jeffrey's improvements in the celloidin method have been 
described in considerable detail by Plowman. 1 Sections of hard 
stems and roots cut by this method could hardly be surpassed, 
and they are perfectly adapted to the requirements of photomicro¬ 
graphy. The following is a brief abstract of Plowman's paper: 

1. Preparation of Material. —-Dead and dry material should be 
repeatedly boiled in water and cooled to remove air. An air-pump 

1 A. B. Plowman, The Celloidin Method with Hard Tissues, Botanical Gazette , 37:456-461, 

1904. 


128 


METHODS IN PLANT HISTOLOGY 


may be used in addition. Living material may be fixed in a mixture 


of picric acid, mercuric chloride, and alcohol: 

Mercuric chloride, saturated solution, in 30 per cent alcohol. 3 parts 

Picric acid saturated solution, in 30 per cent alcohol. 1 part 


Fix 24 hours, and wash by passing through 40, 50, 60, 70, and 80 
per cent alcohol, allowing each to act for 24 hours. 

2. Desifilication, etc. —Silica and other mineral deposits are 
removed by treating with a 10 per cent aqueous solution of com¬ 
mercial hydrofluoric acid. The material is tranferred to this solu¬ 
tion from water or from the 80 per cent alcohol. The process may 
require 3 or 4 days, with one or two changes of the acid and frequent 
shaking of the bottle. An ordinary wide-mouthed bottle, coated 
internally with hard paraffin, should be prepared, since the acid is 
usually sold in bottles with narrow necks. The bottles are easily 
prepared by filling them with hot paraffin and simply pouring the 
paraffin out. Enough will stick to the bottle to protect the glass 
from the acid. Wash in running water 3 or 4 hours. 

3. Dehydration. —Use 30, 50, 70, 90, and 100 per cent alcohol, 
allowing 12 hours in each grade. 

4. Infiltration with Celloidin. —There should be ten grades of 
celloidin: 2, 4, 6, 8, 10, 12, 14, 16, 18, and 20 per cent. Transfer 
from absolute alcohol to the 2 per cent celloidin. (We should prefer 
a previous treatment with ether-alcohol.) The bottle should be 
nearly filled, and the stopper should be clamped or wired in place. 
Put the bottle on its side in a paraffin bath at 50° to 60° C. for 12 to 
18 hours. Cool the bottle quickly in cold water, taking care that 
the water does not get into the bottle. Pour out the 2 per cent 
solution (which, as well as all other solutions, may be used repeat¬ 
edly), and replace it with the 4 per cent, and proceed in the same way 
with the other grades. When the 20 per cent solution is reached, a 
further thickening is gained by adding a few chips of dry celloidin 
from time to time until the mixture is quite stiff and firm. Remove 
each block with the celloidin adhering to it and harden it in chloro¬ 
form for 12 hours. Then transfer to a mixture of equal parts of 
glycerin and 95 per cent alcohol, where the material should remain 
for a few days before cutting. 

Cutting, Staining, and Mounting. —Although 10 ix is usually thin 
enough, sections are readily cut as thin as 5 /jl by this method. 




THE CELLOIDIN METHOD 


129 


Remove the celloidin before staining by treating 10 to 15 minutes 
with ether; then wash in 95 per cent alcohol and transfer to water, 
and then to the stain. Stain to a fairly dense purple in an aqueous 
solution of Erlich’s haematoxylin; wash in dilute aqueous solution of 
calcium or sodium carbonate, and then in two changes of distilled 
water. Add a few drops of alcoholic solution of equal parts of 
Griibler’s alcoholic and aqueous safranin, and stain to a rich red. 
A dilute stain acting 1 to 2 hours is better than a more concentrated 
stain acting for a shorter time. Transfer directly to absolute alcohol, 
clear in xylol, and mount in balsam. 

Haidenhain’s iron-haematoxylin is a very satisfactory stain for 
photographic purposes. 

The celloidin method has its disadvantages as well as its advan¬ 
tages. It is extremely slow and tedious, and it is rarely possible to 
cut sections thinner than 10 n, while, on the other hand, it gives 
smoother sections. 

Succulent tissues, which are usually damaged by the paraffin 
method, are easily handled without any injury in celloidin. The 
fact that the method may be used without heat is often a further 
advantage. Stems and roots which cannot be handled at all in 
paraffin cut well in celloidin, and much larger sections can be cut than 
in paraffin, but most material of this kind can be cut without any 
imbedding. 

When material is to be imbedded, use celloidin as a last resort. 
Use paraffin when you can, celloidin when you must. 


CHAPTER XI 

THE CELLULOSE ACETATE METHOD 

We believe that the cellulose acetate method is destined to 
prove as successful with hard woody tissues as the Venetian turpentine 
method has proved with unicellular and filamentous forms; but, it 
must be confessed that investigators have been unsuccessful in their 
experience with cellulose acetate. Nevertheless, refractory material, 
like oak and even harder woods, has yielded thin transverse sections. 
Cellulose acetate does not injure the finer details of structure, and, 
on that account, will be superior to hydrofluoric acid, when students 
have solved the present uncertainties in regard to it. We have such 
confidence in the value of the method that we are quoting, in full, 
Mrs. Williamson’s short account in the Annals of Botany for January, 
1921, which is the only published information upon the subject: 

A NEW METHOD OF PREPARING SECTIONS OF HARD 
VEGETABLE STRUCTURES 

In order to prepare sections of hard vegetable structures it is essential 
that some method should be devised by which the structure is not only 
embedded but softened, so that sections can be cut easily and smoothly. 
After various methods had been tried, the cellulose acetate method success¬ 
fully used by Dr. Kernot for embedding and sectioning the fabric of aeroplane 
wings was used. It was discovered that this method not only embedded 
hard vegetable structures, but also softened them so that sections are easily 
obtained. It proved best to use cellulose acetate of French manufacture 
made from pure cellulose, as the viscosity is more uniform than that of 
English manufacture, which is obtained from the cellulose of wood. 

In the preliminary experiments pieces of oak and beech, cut into half¬ 
inch cubes, were passed through strengths of alcohol, then placed in pure 
acetone for two hours and finally into a 12 per cent solution of cellulose 
acetate in acetone. There they were left for two months and excellent 
sections were obtained. Further experiments showed that the passage 
through alcohols was unnecessary. In the final experiments the pieces of 
wood were placed in water and the air removed from them, after which they 
were put into pure acetone for 1 to 2 hours and finally into the solution of 
cellulose acetate. It was found that the length of time of immersion in the 

130 


THE CELLULOSE METHOD 


131 


solution of cellulose acetate necessary for softening the tissues varied with 
the hardness of the wood, the minimum time for soft woods being two days; 
for woods such as oak and beech, at least six days are required. Experi¬ 
ments were tried with sal (Shorea robusta ) and Pyingadu (Xylia dolabriformis ), 
one of the Indian ironwoods, which is extremely hard. After fourteen days 
in the cellulose acetate solution it was possible to obtain transverse sections 
of these hard woods. The cellulose acetate solution is therefore capable of 
softening even the hardest wood in a relatively short time. 

In order to stain sections—either hand or microtome—obtained by this 
method, it is necessary to wash them in pure acetone for 1 to 2 minutes to 
remove the cellulose acetate, wash in alcohol 1 to 2 minutes, and pass on to 
the stains selected. Various staining methods for cell walls—such as anilin 
chloride, methylene blue, and Congo red, ammoniacal fuchsin and Kleinen- 
berg’s haematoxylin, etc.—were tried with success. A comparison with 
stained sections of untreated wood revealed no differences. Delicate tissues 
in the wood and hyphae of fungi infecting the wood also stain well and are 
unaffected by the treatment. 

A satisfactory method of preparing sections of hard vegetable structures 
is therefore supplied by the use of a 12 per cent solution of cellulose acetate 
in pure acetone for softening and embedding.—H. S. Williamson, Imperial 
College of Science and Technology. 

Correspondence with Mrs. Williamson indicates that the various 
brands of cellulose acetate behave differently. Cellulose acetate 
obtained from wood is unsatisfactory. We found that cellulose ace¬ 
tate made from photographic films was also unsatisfactory. Mrs. 
Williamson used a cellulose acetate sold by Cellon (Richmond) Ltd., 
22 Cork Street, London, England, and manufactured by the Societe 
Chemique des Usines du Rhone. The time may be shortened by 
keeping the temperature at 40° C. 

In making the solution, use 12 g. of cellulose acetate to 100 c.c. 
of pure acetone. 

It would be worth while to try to adapt this method to the paraffin 
method. 


CHAPTER XII 


SPECIAL METHODS 

In the preceding chapters it has been our object to present the 
principal methods of preparing plant material for microscopic 
examination in such a way that good preparations may be obtained 
as soon as possible. The schedules afford an easy introduction to the 
various methods, but it is hoped that the methods have been so 
presented that the student has become grounded in the fundamentals, 
because some modification is likely to be necessary in nearly every 
piece of research. A few methods designed to meet special difficulties 
are given in this chapter and others are mentioned in the second 
part of the book, in connection with various laboratory types. 

VERY LARGE SECTIONS 

It is sometimes desirable to cut very large sections. Sections 
as large as a cornstalk may be cut freehand or in celloidin. A section 
of a stem of Zamia 5 or 6 cm. in diameter is difficult to handle by the 
usual methods. If a large microtome, such as is used in cutting 
complete sections of large brains, is available, the piece of stem is 
easily held for the cutting. Some of the medium-sized sliding 
microtomes now have a rigid clamp which will grip a block 3 cm. 
square. The lower part of the piece can then be trimmed to fit the 
clamp, leaving the upper part round, so that sections across a stem 
6 or 7 cm. in diameter may be cut without much difficulty. With a 
rather soft stem, like Zamia , the surface must be flooded with 95 per 
cent alcohol after each section, if it is desirable to cut thin sections. 
From stems 3 cm. in diameter, sections can be cut at about 20 to 
30 ji. If the section is not more than 3 or 4 cm in diameter, it can 
be mounted on a 50X75 mm. slide. Sections 6 or 7 cm. in diameter 
can be mounted on lantern slides; if large covers are not available, 
use another lantern slide for a cover. It will be easier to get neat 
mounts if the cover is cut down so as to leave a margin 2 or 3 mm. 
wide. It is not easy to mount a thick section between 2 lantern 
slides of the same size, on account of the balsam which oozes out at 


132 


SPECIAL METHODS 


133 


the edges. Such preparations may be used directly as lantern slides. 
Large sections of the stem of a tree fern make good mounts without 
any staining. Large sections of cornstalk are rather hard to cut, 
because the rigid bundles tear through the soft parenchyma. Flood¬ 
ing with 95 per cent alcohol facilitates the process. A slight harden¬ 
ing is sufficient, so that about 4 or 5 sections can be cut in a minute. 
The corn stem, and similar structures, can be cut in paraffin by Miss 
Langdon’s method, described on page 93. 

STONY TISSUES 

Sections of the stony tissues of hickory nuts, walnuts, peach 
stones, and similar refractory substances cannot be cut by ordinary 
methods. 

With a fine saw, saw sections about 1 mm. in thickness. Rub 
a section between 2 pieces of fine sandpaper until it is not more 
than half a millimeter in thickness. Then rub it between 2 smooth 
hones, keeping the hones wet with water. When the section is thin 
enough, wash it thoroughly in water, using a pipette to rinse off any 
particles of dirt. Dehydrate in absolute alcohol, clear in clove oil, 
and mount in balsam. The long, narrow pores show better without 
any clearing. In this case, dry the section thoroughly, heat a few 
drops of balsam on the slide to drive off the solvent, put the section 
into the balsam, and add a cover. The air caught in the long, narrow 
pores will make them appear as black lines. Sections of most nuts 
show excellent detail without any staining. Thin sections, however, 
may be stained in the usual way. 

PETRIFACTIONS 

Paleobotany has made such rapid progress during the last ten 
years that scarcely any problem involving the anatomy of living 
vascular plants can be investigated intelligently without some knowl¬ 
edge of Mesozoic and Paleozoic forms. Material, especially that of 
Paleozoic Pteridophytes and Gymnosperms, is becoming available, 
and consequently it is increasingly necessary for laboratories to have 
apparatus and technic for cutting rock-sections. 

The outline of the process of cutting a rock-section is very simple: 

1. Saw the rock into two pieces. 

2. Polish the cut surface. 

3. Fasten the cut surface to a piece of glass with hot shellac. 


134 


METHODS IN PLANT HISTOLOGY 


4. With the saw, make another cut, as close to the glass as possible, 
so as to leave a thin section firmly fastened to the glass. 

5. Grind and polish until the section is as thin as possible, or as thin as 
you want it. 

6. Wash all polishing powder off with water. 

7. Dry completely and, either with or without moistening in xylol, mount 
in balsam. 

A word of suggestion in regard to these various points may not 
be amiss. 

1. Most rock-sections are cut with a rather expensive and quite 
complicated instrument, called a “petrotome.” The saw is of the 
circular type, is made of tin or other soft metal, has no teeth, but has 
diamond dust driven into the margin. A rigid clamp holds the 
object, and the saw, constantly cooled by a stream of water, gradually 
cuts through the specimen. If the piece to be cut is more than 5 
or 6 cm. in diameter, a band saw is better; and if the piece is 10 or 
20 cm: in diameter, the band saw is necessary. 

2. The cut surface is most easily polished on a revolving brass 
plate, kept wet and liberally powdered with fine carborundum. 
When the surface has become even and smooth, the specimen is 
ready for the next step. 

3. Fasten the polished surface to the glass slide upon which the 
section is to be mounted. Plate glass 3 or 4 mm. thick is best for 
sections larger than 3 or 4 mm. square. Gradually heat the slide 
until it is quite hot. Melt upon the slide the thin flakes of white 
shellac used by painters; heat the object and press the polished 
surface very firmly into the melted shellac. Canada balsam, from 
which the xylol has been driven off by heating, can be used instead 
of shellac. Much of the Paleozoic material is in the form of coal 
balls. After the ball has been cut in two, it is often difficult to hold 
the hemispherical piece in a clamp, especially if the piece is small. 
In such cases, it is better to fasten the polished surface to a convenient 
piece of marble, about 2.5 cm. thick, and 5 or 6 cm. square. The 
marble is easily held in the clamp. As soon as the slide, or marble, 
and object are cool, the next cut can be made. 

4. Fasten the object in the clamp and saw as close to the glass, 
or marble, as possible, thus leaving a thin section cemented to the 
slide or marble. If marble has been used, the section is removed 
by heating or by dissolving it off with xylol. It can then be fastened 


SPECIAL METHODS 


135 


to the glass slide for grinding and polishing. Anyone who can handle 
tools should soon be able to cut a section 1 mm. thick. A skilled 
technician can cut sections as thin as 0.5 mm. 

5. The second grinding must be very careful and accurate. Do 
the polishing on the revolving disk. The glass slide allows one to 
note how the process is progressing. 

6. When the section becomes thin enough, or even before if it 
begins to crack, wash off the powder. 

7. It is usually a good plan to use rather thick balsam for mount¬ 
ing, even if it should be necessary to heat it a little to make it flow well. 

By this method, sections of fossils 10X15 cm. have been cut 
thin enough for examination with a 4 mm. objective. Sections 
3 or 4 mm. square have been cut thin enough for satisfactory examina¬ 
tion with a 2 mm. oil immersion lens. Figure 26 shows that a reliable 
study can be made from sections cut from the solid rock. 

Of course, this method can be used for such objects as walnut 
and hickory shells. 

THICK SECTIONS 

It is sometimes desirable to make very thick sections to show 
general topography rather than detail. A longitudinal section of 
the fully grown ovule of Ginkgo or a cycad may be cut as thick as 
3 to 5 mm. so as to include the entire group of archegonia. A slab 
can be cut from each side of the ovule with a fine saw, and a razor 
can be used for smoothing. If the section is from fresh material it 
should be fixed, washed, etc., with about the same periods as if it 
were to be imbedded in paraffin. When thoroughly cleared in xylol, 
the section should be put into a flat museum jar of suitable size and 
kept in xylol. Even before the stony coat of a cycad becomes too 
hard to be cut with a razor such thick sections are very instructive. 
Stain very lightly, or not at all. Sections of Zamia or other cycad 
stems, 2 mm., or even 5 mm. thick, make instructive mounts, since 
they show the peculiar course of the bundles, a feature which is 
largely lost in thin sections. 

Krause prepared large objects very effectively by dehydrating, 
clearing in xylol, and then transferring to cedar oil. Sections of an 
apple, either longitudinal or transverse, about 3 or 4 mm. thick, 
cleared in this way, are very instructive. Strawberries, gooseberries, 
and similar objects treated in this way afford a kind of study which 
is too often neglected. 


136 


METHODS IN PLANT HISTOLOGY 


Miss Langdon cleared 15-mm. cubes of Dioon stem in a similar 
way, but used equal parts of xylol and carbon disulphide instead of 
cedar oil. The course of the bundles could be traced perfectly, 



Fig. 26 .—Sphenophyllum pleurifoliatum: transverse section of a stem, cut from a coal ball 
of the Upper Carboniferous. Eastman Commercial Ortho film, Wratten E (orange) filter, J. 
Swift and Son 1-inch lens; arc light; exposure, 1 second. Slide by Lomax, negative by Sedgwick. 
X19. 





















SPECIAL METHODS 


137 


LAND’S GELATIN METHOD 

It is sometimes desirable to get sections of partly disorganized 
material. A matrix is necessary to hold the parts in place, but 
dehydration may make the tissue unnecessarily hard to cut. 

Soak ordinary gelatin (which can be obtained at the grocery) in 
water until no more is taken up. Then drain off the excess water 
and liquefy the gelatin by heating. Place the material—previ¬ 
ously soaked in water—in the melted gelatin and keep it there 
for several hours. Place also in the gelatin some small blocks of 
hard wood to serve as supports in the microtome. The material 
to be sectioned is oriented in a gelatin matrix on the supporting 
blocks, cooled until the gelatin sets, and then placed in strong 
formalin to harden the gelatin. In cutting, flood the knife with 
water. 

If the material is to be stained, stain it in bulk before putting it 
into the gelatin, since the gelatin stains very deeply. Of course, 
the gelatin could be dissolved with hot water, or hot water and acetic 
acid, but all the advantage of a matrix would be lost. 

It would be worth while to try this method thoroughly with soft, 
succulent tissues and with hard tissues which become still harder 
if dehydrated. 

SCHULTZE’S MACERATION METHOD 

Various solutions are used to separate a tissue into its individual 
cells. These solutions dissolve or weaken the middle lamella so that 
the cells are easily shaken or teased apart. Schultze used strong nitric 
acid and potassium chlorate. Put the material, which should be in 
very small pieces, into a test-tube; pour on just enough nitric acid 
to cover it, and then add a few crystals of potassium chlorate. Heat 
gently until bubbles are evolved, and let the reagent act until the 
material becomes white. Four or five minutes should be sufficient. 
The fumes are disagreeable and are very injurious to microscopes. 
Pour the contents of the tube into a dish of water. After the material 
is thoroughly washed in water, it may be teased with needles and 
mounted, or it may be put into a bottle of water and shaken until 
many of the cells become dissociated. 

After a thorough washing in water, the material may be stained. 
The large tracheids of ferns, dissociated in this way and stained in 
safranin or methyl green, make beautiful preparations. 


138 


METHODS IN PLANT HISTOLOGY 


JEFFREY’S MACERATION METHOD 

A method which I saw at Toronto and which gives much better 
results was credited to Professor Jeffrey. Wood is cut or split into 
sections about 300 ju thick, which are then boiled and cooled until 
free from air. The macerating fluid consists of equal parts of 10 per 
cent nitric acid and 10 per cent chromic acid. The time will vary 
with different woods, but is likely to be about 24 to 48 hours if the 
temperature is about 35° C. When properly macerated the cells may 
be shaken apart or are very easily teased apart. Before staining, 
the material should be very thoroughly washed to remove the acid. 
A study of such material is very valuable in modern anatomical work. 

Maceration methods which act in a few minutes are likely to be 
so violent that fine details will not be preserved. 

PROTOPLASMIC CONNECTIONS 

As a rule, protoplasmic connections are not likely to be seen in an 
ordinary preparation. It used to be thought that the rather large 
protoplasmic strands seen at the sieve plates of the pumpkin and other 
Cucurbitaceae were exaggerated examples of protoplasmic continuity; 
but, as a matter of fact, the large strands do not extend entirely 
through the plate. The real continuity, through the middle lamella, 
is scanty and hard to demonstrate. 

Very satisfactory material for the demonstration of the connecting 
strands is furnished by the seeds of the Japanese persimmon, Diospiros 
Kaki. Fix in formalin alcohol (10 c.c. formalin to 50 c.c. of 70 per cent 
alcohol) or in a mixture of equal parts 70 per cent alcohol and glycerin 
(Fig. 27). 

Good views of the strands are most abundant in sections cut 
parallel with the flat surface of the seed. No imbedding is either 
necessary or desirable. Clamp the seed in the microtome directly, 
or fasten it to a wooden block with cellulose acetate (easily made by 
dissolving a photographic film in acetone; of course the emulsion 
should be removed), with celloidin, or even with glue. Cellulose ace¬ 
tate seems to be the best. Cut sections 8 to 12 p thick; place them 
in ether several hours to remove any fatty substances; remove the 
ether with absolute alcohol; transfer to 95 per cent alcohol, then to 
50 per cent alcohol, and then to water. Stain in iron-alum haema- 
toxylin. The following schedule for Diospiros Kaki will introduce 
the method. 


SPECIAL METHODS 


139 


1. Fix in formalin alcohol or in glycerin alcohol 4 or 5 days. 

2. Cut sections 8 to 12 /x thick. 

3. Ether, several hours. 

4. Absolute alcohol, 95 per cent and 50 per cent alcohol, 10 minutes each. 

5. Wash in water, 5 minutes. 

6. Iron alum, 4 per cent, 8 to 24 hours. 

7. Wash in water, 30 minutes. 

8. Stain in § per cent haematoxylin 24 hours. 

9. Wash in water. At this stage, examine carefully, because it may not 
be necessary to reduce the stain in iron-alum. If the connections are 
not too deeply stained, simply dehydrate, clear, and mount in balsam. 

After the first 5 stages, a strong stain in DelafiekTs haematoxylin, 
10 to 24 hours, followed by a very weak hydrochloric acid, has given 



Fig. 27 .—Diospiros Kaki: section of endosperm prepared as described for this species. X780 

good results. A sharp stain in crystal violet, differentiated with 
orange in clove oil, often fails, but sometimes succeeds; and, when 
successful, the connections stand out beautifully. 

The endosperm of Phytelephas (vegetable ivory), of dates and 
many other palms, and probably most hard endosperms, will show 
the connections by the methods just described; but in many cases 
it is necessary to resort to special methods in order to demonstrate 
the continuity. In these special methods a reagent is used which 
causes the membranes to swell before the stain is applied. It is only by 
such an exaggeration that the more delicate connections can be shown. 











140 


METHODS IN PLANT HISTOLOGY 


Put thin sections of fresh material into a mixture of equal parts 
of sulphuric acid and water; and allow the reagent to act for 2 to 
10 seconds. Wash the acid out thoroughly in water and stain in 
anilin blue. According to Gardiner, this stain should be made by 
adding 1 g. of the dry stain to 100 c.c. of a saturated solution of 
picric acid in 50 per cent alcohol. The staining solution is then 
washed out in water, and the sections are mounted in glycerin. The 
sections may be dehydrated, cleared in clove oil, and mounted in 
balsam. The anilin blue may be used in 50 per cent alcohol acidu¬ 
lated with a few drops of acetic acid. 

Chloro'iodide of zinc may be used instead of sulphuric acid. Treat 
the fresh sections for 2 hours with the iodine and potassium-iodide 
solution used in testing for starch; then treat about 12 hours with 
chloroiodide of zinc. Wash in water and stain in anilin blue. 
Examine in glycerin. 

Meyer’s pyoktanin method is one of the best. The reagents are 
as follows: 

1. Iodine, potassium iodide solution: iodine 1 part, potassium iodide 
1 part, water 200 parts. 

2. Sulphuric acid 1 part, water 3 parts; this mixture to be saturated 
with iodine. 

3. Pyoktanin coeruleum 1 g., water 30 c.c. This pyoktanin is a very 
pure methyl violet obtained from E. Merck in Darmstadt. 

Put sections of the date seed into a watch glass full of the first 
solution, and allow it to act for a few minutes; then mount in a drop 
of the solution. The connections will be only very faintly stained, 
showing a slightly yellowish color. At the edge of the cover, add a 
drop of the second solution. The preparation will darken a little. 
Then allow a small drop of the third solution to run under the cover. 
Allow the stain to act for about 3 minutes. Then plunge the whole 
preparation into water. The action should be stopped before 
the entire section has become blue. Now wash the section quickly. 
If there are annoying, granular precipitates, remove them with 
a soft brush. Mount in glycerin. The membrane should be a 
clear blue, while the protoplast and connections should be a blue 
black. 

The following is Strasburger’s modification of Meyer’s method, 
and shows the connections with great distinctness: 


SPECIAL METHODS 


141 


Permanent preparations may be secured by the following method: 

1. Treat the fresh sections with 1 per cent osmic acid, 5 to 7 minutes. 

2. Wash in water 5 to 10 minutes. 

3. Treat with a solution of iodine in potassium iodide (0.2 per cent 
iodine and 1.64 per cent potassium iodide), 20 to 30 minutes. 

4. Transfer to 25 per cent sulphuric acid, which should act for at least 
half an hour; 24 hours may be necessary. 

5. Bring the sections into 25 per cent sulphuric acid which has been satu¬ 
rated with iodine. Add a drop of Meyer’s pyoktanin solution (1 g. pyok- 
tanin coeruleum as sold by E. Merck in Darmstadt in 30 c.c. of water). 

In about 5 minutes the sections will be stained sufficiently and 
can be examined in glycerin. If there are annoying precipitates, 
remove them with a soft brush. 

According to Meyer, the swelling is an advantage only when 
the walls are very thin. When the walls are thick, the connections 
show better without any previous swelling. 

Try the following method with the seeds of Diospiros, Latania 
Chamerops, Phoenix, or Phytelephas: Soak in water and cut thin 
sections. Extract the oily and fatty substances with xylol; wash 
in 95 per cent, or in absolute alcohol; stain in anilin blue (Hoffman’s 
blue 1 g. dissolved in 150 c.c. of 50 per cent alcohol) for a few minutes. 
Examine in glycerin. This method succeeds very well with seeds of 
the date, which is sold at all groceries. 

Permanent preparations may be secured by the following method: 

1. Fix in 1 per cent osmic acid, or in absolute alcohol, 5 to 10 minutes. 

2. Stain for 24 hours in Delafield’s haematoxylin. 

3. Wash for a few minutes in acid alcohol (5 drops of hydrochloric acid 
in 50 c.c. of 70 per cent alcohol). 

4. Wash for a few minutes in ammonia alcohol (5 drops of ammonia to 
50 c.c. of 70 per cent alcohol). 

5. Dehydrate in absolute alcohol, clear in xylol, and mount in balsam. 

STAINING CILIA 

The cilia of the large spermatogoid of Ginkgo and the cycads 
stain beautifully in iron-alum haematoxylin, which not only stains 
the cilia but even differentiates the free portion from the part between 
the blepharoplast and the surface. 

The cilia of sperms of Bryophytes and Pteridophytes stain better 
with gentian-violet or crystal-violet. The periods are long; not 
less than 30 minutes, and often several hours will be required. 


142 


METHODS IN PLANT HISTOLOGY 


The cilia of the motile spores of Thallophytes may often be 
demonstrated by allowing a drop of the iodine solution used in testing 
for starch to run under the cover. 

Zimmermann gives the following method: Bring the objects into 
a drop of water on the slide and invert the drop over the fumes of 
1 per cent osmic acid for 5 minutes. Allow the drop to dry. Then add 
a drop of 20 per cent aqueous solution of tannin, and after 5 minutes 
wash it off with water. Stain in a strong aqueous solution of fuchsin 
(or carbol fuchsin) for 5 minutes. Allow the preparation to dry 
completely, and then add a drop of balsam and a cover. The cilia 
should take a bright red. 

Zimmermann also found the following method satisfactory for 
the cilia of the zoospores of algae and fungi: Fix by adding a few 
drops of 1 per cent osmic acid to the water containing the zoospores; 
then add an equal amount of a mixture of fuchsin and methyl violet. 
The fuchsin and methyl violet should be 1 per cent solutions in 95 per 
cent alcohol. In a few seconds the cilia stain a bright red. 

CHONDRIOSOMES 

During the past fifteen years the terms chondriosomes, mito¬ 
chondria, Chondriokonten, Chondromiten , etc., have become increas- 



Fig. 28.—Cells from the periblem of the root-tip of Allium cepa: A, chondriosomes; B, 
canaliculi: fixed in Bensley’s solution and stained in iron-alum haematoxylin. X 1,200. 

ingly frequent in botanical literature. These “ chondriosomes,” 
as we shall call them, are minute structures, probably present in most 
cells, but not differentiated by the most usual methods and generally 
overlooked when they might be seen. Most of them are as small as 
bacteria and bear a superficial resemblance to coccus, spirillum, 
and bacillus forms (Fig. 28A). 






SPECIAL METHODS 


143 


Many fixing agents either destroy the mitochondria or make it 
almost impossible to demonstrate them. Fixing agents containing 
alcohol or any considerable percentage of acid are to be avoided. 
Benda’s solution, followed by Haidenhain’s iron-alum haematoxylin, 
will give good results. A solution recommended by Bensley is good 
also for plant material. 


Bensley’s Solution.— 

Osmic acid 2 per cent. 1 part 

Corrosive sublimate (HgCl 2 ) per cent. 4 parts ' 


Add 1 drop of glacial acetic acid to 10 c.c. of this solution. Fix 
for 24 to 48 hours and wash thoroughly in water. On the slide, 
bleach with hydrogen peroxide; wash in water; treat with the iodine 
solution used in testing for starch; then wash in water. The slide 
is now ready for staining. We recommend the usual Haidenhain’s 
iron-alum haematoxylin. 

Bensley recommends the following method which we have found 
rather uncertain, but which, when successful, yields magnificent 
preparationsOn the slide, bleach for 2 or 3 seconds in a 1 per cent 
aqueous solution of permanganate of potash; then treat with a 
5 per cent aqueous solution of oxalic acid until the preparation 
becomes white (a few seconds); wash in water, and then stain as 
follows: 

1. Copper acetate (neutral) saturated solution in water, 5 to 10 minutes. 

2. Wash in water. 

3. ^ per cent haematoxylin, 5 to 10 minutes. 

4. Wash in water. 

5. Potassium bichromate (neutral) 5 per cent solution in water until the 
preparation blackens, usually 30 seconds or less. 

6. Differentiate in Weigert’s ferricyanide solution. 


Borax. 2.0 g. 

Ferricyanide of potassium. 2.5 g. 

Water. 200.0 c.c. 


7. Wash in water and proceed as usual. 

CANALICULI 

By using special methods, Bensley has obtained views of the 
protoplasm of plants, quite different from those seen in ordinary 
preparations. In the cell of a root-tip a series of small canals, or 
vacuoles, appears, which is much more definite and extensive than 







144 


METHODS IN PLANT HISTOLOGY 


the usual display of vacuoles and which appears before any vacuoles 
can be recognized in preparations made in the usual way (Fig. 28 B). 
Being a zoologist, he has called these vacuoles “canaliculi.” 

VASCULAR BUNDLES IN LIVING TISSUES 

In studying venation, and in tracing the course of vascular 
bundles in large ovules and in other organs, it is often an advantage 
to use a stain. If a stem of Impatiens be cut under water, and the 
cut surface be then placed in a dilute aqueous solution of eosin, the 
eosin will rise in the vessels, making them very prominent. The 
outer bundles of the large ovules of cycads are very easily studied 
by this method. The inner bundles also may be seen by opening 
the seed and removing the endosperm. 

If such preparations could only be cleared, they would be still 
more valuable, but the effect is due to the presence of the staining 
fluid in the vessels, and any subsequent treatment diffuses or destroys 
the stain. Perhaps a little experimenting might obviate the difficulty. 

STAINING LIVING STRUCTURES 

Some stains will stain living structures. Cyanin, methyl blue, 
and Bismarck brown have been recommended for this purpose. The 
solutions should be very dilute, not stronger than 1:10,000 or 
1:500,000. The solutions should be very slightly alkaline, never 
acid. It is claimed that such solutions never stain the nucleus, and 
that if the nucleus stains at all, it is an indication that death is taking 
place. 

Campbell succeeded in staining the living nuclei in the stamen 
hairs of Tradescantia by using dilute solutions of dahlia and of 
methyl violet (0.001 to 0.002 per cent in water). Dividing nuclei 
were stained. 

For determining the stage of development of fresh material it is 
often necessary to use a stain. For this purpose stronger stains may 
be used, since it is unimportant whether the tissue is killed or not. 
An aqueous solution of methyl green or eosin can be recommended. 
With 1 per cent solutions, diluted one-half with water, mitotic 
figures can be recognized with ease. 


CHAPTER XIII 

PHOTOMICROGRAPHS AND LANTERN SLIDES 

While a work like the present book is hardly the place for any 
extended treatment of photomicrography or the making of lantern 
slides, a few simple directions will help the beginner and enable him 
to prepare most of the photomicrographs and lantern slides which 
may be necessary in the classroom. It is assumed that the student 
knows how to handle an ordinary camera and knows how to do his 
own developing. 

In the preparation of this chapter, I am indebted to Dr. Paul J. 
Sedgwick for suggestions, criticism, and considerable revision. 

PHOTOMICROGRAPHS 

For a simple beginning, no apparatus is needed except an ordi¬ 
nary camera and a microscope. Try low powers first and proceed 
gradually to the higher magnifications. Remove both front and 
back lenses from the camera, leaving the lens barrel and the shutter; 
also, remove the eyepiece from the microscope. Bend the micro¬ 
scope to the horizontal position and place the lens of the camera 
close to the ocular end of the microscope and shut out all light at 
this point by winding black cloth around the end of the microscope 
and the barrel of the camera lens. Take great care to have a per¬ 
fectly straight optical axis through the microscope and camera. 

While the camera and microscope can be adjusted so as to secure 
a perfect optical axis by simply putting both instruments on the table 
and raising one or the other—according to the size of the camera—• 
by placing a board under it, such an adjustment is extremely unsatis¬ 
factory, since the least jar may disturb it, and inserting the plate- 
holder is almost sure to disarrange something. It will save time 
if you prepare a board to keep both instruments in position. Select 
a clear board 1 inch thick, about 1 foot wide, and 5 feet long. On 
the top of this board, screw two pieces f inch thick, 1| inches wide, 
and 5 feet long, so as to form a guideway for the camera (Fig. 29 A). 
If the camera is so small that it must be raised to bring it into the 


145 


146 METHODS IN PLANT HISTOLOGY 

optical axis of the microscope, fit to the guideway a board of the 
necessary thickness, and fasten the camera to this board. It is 
absolutely necessary that the preparation to be photographed and 
the ground glass of the camera should be perfectly parallel. The 
board will save time in securing this parallelism. Cut through the 
board a slot l inch wide and extending to within 6 inches of each end. 
By this means the camera can be clamped with the screw used to 

fasten it to a tripod. Also, a 
piece of metal or hard wood 
may be placed over the horse¬ 
shoe base of the microscope 
and with a bolt, preferably one 
with a butterfly nut, the micro¬ 
scope may be held firmly in 
place. This board, with the 
long slot, will be useful in 
making lantern slides. 

As an illuminant, direct 
sunlight, diffuse daylight, a 
gas-mantle lamp, an acetylene 
lamp, a Mazda bulb, an arc 
light, or any strong light may 
be used. Remove the mirror 
from the microscope and allow 
the light to come directly into 
the optical axis. This mirror 
will not be needed in any pho¬ 
tomicrographic work. 

Let us suppose that we are to make a photomicrograph of a 
vascular bundle and that we are using a 16-mm. objective. If 
only a part of the bundle is shown on the ground glass, remove 
the ocular of the microscope. If the illumination is very uneven 
and shows a “flare spot/’ look at the inside of the tube of the micro¬ 
scope. Probably it was not blackened and the “flare spot” was 
due to reflections. Obviate the difficulty by putting a piece of black 
paper inside the tube. Any modern microscope should have the 
tube well blackened inside. Move the light back and forth and 
sidewise to get the best illumination. An easy way to determine 
whether the light is centered is to close down the iris diaphragm below 



A 



^^ 



l-Z 1 L 



Fig. 29.— A, board for photomicrographic and 
lantern-slide work; B, end view with clips to hold 
negatives; C, side view of block to be used on board 
when making lantern slides. 





















PHOTOMICROGRAPHS AND LANTERN SLIDES 


147 


the condenser until the lighted circle on the ground glass is very small. 
Then shift the light or the condenser, for it is possible that the con¬ 
denser is also off center, or both until the illuminated circle is 
centered. After the light source, the condenser, and the objective 
have been brought into line, rack the condenser back and forth until 
the beam of light is focused on the object to be photographed. 
Then make certain that the cone of light at the point where it enters 
the objective has a diameter great enough to fill the aperture of 
the objective. The ordinary form of Abbe condenser is not likely 
to be satisfactory with objectives of 16-mm. focus, and should not be 
used at all with objectives of longer focus. 

Focus the object upon the ground glass. Even with a 16-mm. 
objective, the ordinary ground glass is rather coarse for accurate 
focusing. Always examine the image with a focusing lens. A 
brilliant view may be obtained by fastening a thin cover-glass to the 
ground glass with a small drop of balsam. At this spot the image 
may be examined very critically. Of course, as in any photography, 
the ground side of the glass should be nearest the object, occupying 
exactly the place which is to be occupied by the emulsion side of the 
plate. Do not focus indiscriminately, but be sure that the image 
is sharp at the level of the ground side of the glass. It is a good plan 
to make a cross upon the ground glass with a pencil or pen, and then 
add a drop of balsam and a cover-glass. Focus on this mark and 
fix the focusing glass at this level. The cheap tripod lenses are good 
for this purpose. 

The time of exposure will vary with the magnification, the 
intensity of the light, and the speed of the plate. The exposures will 
be much longer than in ordinary photography. It is better to use 
artificial light, since one can more quickly learn to estimate the 
length of exposure when the intensity of the light is constant. A 
slow plate, even the very slow contrast plate, is likely to prove most 
satisfactory for the beginner. With a Welsbach lamp, a contrast 
plate of the same speed as a lantern slide, and a 16-mm. objective 
used without an ocular, or Abbe condenser, try an exposure of 30 
seconds. Develop the negative in whatever solution is recommended 
in the directions which come with every box of plates. If the negative 
is too weak, make a longer exposure; if too dense, shorten the 
exposure. A little experience with your apparatus will soon enable 
you to estimate the length of exposure with some certainty. We 


148 


METHODS IN PLANT HISTOLOGY 


frequently use lantern-slide plates for tests and for small photo¬ 
micrographs. The Cramer lantern slides and contrast plates of 
larger sizes have the same speed and, consequently, one can determine 
the length of exposure by using a cheap lantern slide. In making 
tests, it will save both time and money to expose for 5 seconds and 
then push in the dark slide so as to cover a part of the plate; then 
expose 5 seconds longer and push the slide in a little farther, etc. 
In this way you can make 4 or 5 exposures on a lantern-slide plate 
showing exposures of 5, 10, 15, 20, and 25 seconds, the first exposure 
being 5 seconds and the last, 25 seconds. By making several expo¬ 
sures on 1 plate, with the first probably underexposed and the last 
probably overexposed, you can note which exposure is best and will 
waste only 1 plate in getting the time. A print from such a negative 
is valuable, since it enables one to judge very accurately the printing 
quality of the various exposures. 

A slow contrast plate such as mentioned above will prove satis¬ 
factory for photographing some subjects and because of ease in 
manipulation it offers certain advantages for the beginner; but it 
is not the most satisfactory in a majority of cases. It is necessary 
to use a plate which is color corrected when one is using ray filters 
to increase or decrease the contrast between differently stained 
portions of a preparation or to secure greater detail than could be 
obtained without a filter. We have used the Cramer Iso plates and 
the Eastman Standard Orthonon plates a great deal. They are 
orthochromatic, sensitive to a large part of the spectrum, and can be 
used with the various filters, with the exception of the reds. When 
it is necessary to use a red filter, we suggest the Wratten Panchromatic 
plates. 

Professional films are rapidly replacing plates for many phases 
of photographic work. The Eastman Commercial Ortho film was 
used in the making of all the photomicrographs in this edition credited 
to Dr. Sedgwick. This Commercial Ortho film is thick, rather slow, 
and gives a combination of detail and contrast which is very desirable. 
It is heavy enough to lie flat in the Professional Film holders made for 
Professional films. It is strongly orthochromatic and may be used 
with all the filters commonly employed in photomicrography except 
for the reds. If it is necessary to use a red filter in order to make a 
red-stained portion of a preparation appear light, the Eastman 
Commercial Panchromatic film may be used. This film works as 


PHOTOMICROGRAPHS AND LANTERN SLIDES 


149 


well as the Commercial Ortho, has the additional advantage of being 
sensitive to red, but is more difficult to handle because it is fogged 
by the usual red darkroom lamps. Panchromatic films, like panchro¬ 
matic plates, should be developed in an unlighted darkroom, or if 
it seems necessary occasionally to examine the negative during 
development, the special Wratten safelight made for this purpose 
should be employed. The Eastman Kodak Company publishes 
an interesting booklet entitled “Professional Film” which they 
distribute free of charge. 

With fast plates and without filters, a strong light will allow 
exposures of a small fraction of a second but we have had no success 
under such conditions. The ordinary yellow filters used in out-of- 
door work will be worth trying, but if one is contemplating doing 
any considerable amount of serious work we would suggest procuring 
a set of filters made up especially for photomicrography. The set of 
9 filters known as the Wratten “M” filters made by the Eastman 
Kodak Company is very useful. With these filters used singly and 
in combination it is possible to utilize selected portions of the spectrum 
and thus to increase or decrease the contrast either between the 
specimen and background or between the variously stained portions 
of the specimen. For the beginner, perhaps the most confusing part 
of the procedure is in choosing the proper filter. The yellow and 
orange filters will probably be the ones most frequently used, for 
when they are used with a good orthochromatic or panchromatic 
plate or film the result will be an approximately correct rendering of 
the color values. Some workers rarely use any filters except the yellow 
and orange but very frequently other filters will give better results. 

It is not possible to state just which filters will be best for each 
set of conditions but certain rules will aid in determining. To obtain 
the greatest amount of contrast between the specimen and background 
use a filter or combination of filters which transmits only that portion 
of the spectrum completely absorbed by the specimen. If the result 
is unsatisfactory because detail within the specimen has been sacri¬ 
ficed in favor of contrast between the background and specimen, try 
a filter or combination of filters whose spectral transmission is not 
exactly limited to the part of the spectrum absorbed by the object. 
To eliminate contrast between the object and its background use a 
filter or a combination of filters transmitting that portion of the 
spectrum also transmitted by the object. After a little experience 


150 


METHODS IN PLANT HISTOLOGY 


one will be able to estimate the probable photographic result by 
making a visual examination with first one and then another of the 
filters in place. 

An example of how the filters were chosen in making one particular 
photomicrograph of a stem section may be suggestive. The specimen 
was well stained in safranin and anilin blue. If the photograph had 
been made on an ordinary uncorrected plate and without a filter the 
result would have been unsatisfactory. The red of the safranin 
would have taken as pure black while the parts of the specimen stained 
in anilin blue would hardly appear in the photograph because the 
plate was not sensitive to red and very much oversensitive to blue 
giving the blue almost the value of white light. Therefore we used 
a color-corrected plate and a combination of the Wratten “B” and 
“E” filters which gave the maximum contrast for the anilin blue since 
the spectral transmission band of the combination is entirely included 
within the limits of the absorption band of the anilin blue. While 
these filters did not give the maximum contrast for the safranin- 
stained portion of the specimen, the contrast was sufficient and the 
result was more satisfactory than if the safranin stain had photo¬ 
graphed as pure black because all detail in the vessel walls would 
then have been lost. 

A booklet, “Photomicrography,” published by the Eastman 
Kodak Company, includes a table showing the spectral absorption 
bands of some of the stains used in botanical microtechnic. It also 
gives data on the transmission spectra of the Wratten “M” filters. 

The Abbe condenser, which should not be used at all with low 
powers, is very useful with objectives of 8-mm. focus and all higher 
powers, especially if the condenser is achromatic. If the condenser 
is not achromatic, it is sometimes a good plan to remove it and 
in its place put a 16-mm. objective, or, for very high powers, even 
an 8-mm. objective. The condenser may be fastened into the con¬ 
denser sleeve by an improvised ring or collar. Zeiss makes a collar 
for this purpose. Spencer microscopes also have such collars. 

In addition to the Abbe condenser, there should be another, 
placed between the microscope and the light. For this purpose, 
the large condenser from a projection lantern may be used. For 
magnifications higher than 100 diameters still another condenser will 
be useful. Place it between the last-named condenser and the 
microscope. 


PHOTOMICROGRAPHS AND LANTERN SLIDES 


151 


With so many condensers, the heat may damage preparations: 
so place between the last-named condenser and the one next the 
light a cooler filled with water or a solution of alum. 

With all these accessories, an additional iris diaphragm is desir¬ 
able. Place it between the middle one of the three condensers and 
the miscroscope, but quite close to the middle condenser. To make 
an efficient adjustment of all these parts requires patience, practice, 
and judgment. 

It will save time and patience if the position of the object to be 
photographed be marked in ink on the slide by vertical and horizontal 
lines, or by a circle around it. Even with these lines, it is none 
too easy to get the object into the desired position on the ground 
glass. Remove the ground glass and let the image fall on a piece 
of white cardboard a short distance back of the camera. If the 
curtains are pulled down, the position of the object in the field and 
the focusing of the condensers will be comparatively easy. 

The desirability of a rigid, straight, and accurate optical bed 
will soon be realized. If one is intending to do much photomicro¬ 
graphic work, the heavy, graduated optical bed is almost a necessity. 
However, if time is no object and patience is abundant, good photo¬ 
micrographs at a magnification of over 1,000 diameters can be made 
with no apparatus except a good camera, a good microscope, and a 
good lamp. 

The relative positions of the various parts, as we have used them 
in making the illustrations for this book, are indicated below: 


Tb 

73 

§ 

o 

o 

=3 

a 

03 

& 

03 

03 

Pi 

O 

C3 

m 

2 

C3 

g 

73 

P 

o 

O 

% 

rO +3 

i) & 
a 1 

S3 

■4-^ £h 

2 

o 

c3 

o 

1 

H 

42 fP 
<1 

S Q O 


73 
O 3 
o o 
O O 


Some data which may be helpful will be found in the legends under 
some of the photomicrographs. 

Easily available and inexpensive help will be found in three 
booklets published by the Eastman Kodak Company, of Rochester, 
New York: “ Photomicrography”; “ Wratten Light Filters ”; “ The 
Photography of Colored Objects. ,, 


152 


METHODS IN PLANT HISTOLOGY 


After a little practice the student will read with profit the more 
extended works on this subject, among which are the following: 
The A, B, C of Photomicrography , by W. H. Walmsley (Tennant & 
Ward, New York); Photomicrography , edited by J. Spitta (Scientific 
Press, London, England); Lehrbuch der Microphotographie , by Dr. 
Richard Neuhaus (Harold Bruhn, Brunswick, Germany). 

LANTERN SLIDES 

Lantern slides are now so universally used in the lecture-room 
that every teacher should be able to make them. Three general 
classes of lantern slides, as far as the technic of making them is 
concerned, will be described: (1) lantern slides by contact, (2) by 
reducing or enlarging, and (3) by copying illustrations. 

1. Lantern Slides by Contact.—This method is very simple. 
Imagine that the lantern-slide plate is a piece of printing-out paper, 
and proceed just as in making a print on paper. Remember that 
dust on the negative or plate causes spots in the print, and that spots 
so small as to be almost unnoticeable in an ordinary print will be 
greatly magnified when they appear on the screen. Brush both 
negative and plate very gently with a soft clean brush before making 
the print. If the negative is 3jX4| inches, it can be placed in a 
printing frame of that size, and the lantern slide placed upon it with 
the two films in contact, just as in printing paper. If there is no 
small printing frame, use a 4X5, a 5X7, or even an 8X10 frame. 
In such cases, put in a piece of clean glass free from scratches or 
bubbles, and lay the negative upon it. Lantern slides may be 
printed from a portion of a 4X5 or some larger negative by simply 
placing the lantern-slide plate over the desired spot. Take great 
care not to scratch the negative. 

A much more satisfactory method is to use a box 10 inches 
square and 16 inches high, inside measurement. The box should be 
painted white inside, and the top should be a strong piece of glass; 
but there should be also a wooden lid, hinged at the back and covered 
inside with some thick, soft cloth. Inside the box, about 4 inches 
from the bottom, place two electric bulbs, one red and the other 
white. To diffuse and tone down the light, a piece of glass, ground 
on one side should be placed a little more than halfway between the 
light and the glass top. If the light is still too strong, place a piece 
of white paper above the ground glass. It is most convenient to 


PHOTOMICROGRAPHS AND LANTERN SLIDES 


153 


have the white light operated by a switch. With a 40-watt light, 
exposures will vary from 1 to 30 seconds, as the negatives are very 
thin or very dense. With properly exposed negatives, the exposures 
will vary 2 to 5 seconds. 

If you are printing in a printing frame with an average negative 
at a distance of 3 feet from a gas-mantle lamp, try an exposure of 
3 seconds; if the negative is weak, shorten the exposure; if strong, 
lengthen it. If you always use the same light at the same distance, 
you should soon be able to estimate the exposures for negatives of 
various densities. 

If a negative is uneven, the distance from the light may be 
increased so as to lengthen the exposure to several seconds, thus giving 
time to shade the weak parts, just as in case of prints on paper. 
If a negative is harsh and shows too much contrast, hold it closer to 
the light and shorten the exposure; if weak and lacking in contrast, 
hold it farther away and increase the time of exposure. 

Be careful not to underdevelop. A lantern slide looks stronger 
in the developer, and even in the hypo, than it really is. 

2. Reducing and Enlarging.—If a slide is to be made from a 4X5 
or larger negative, there must be a reduction. A camera is necessary. 
A 3i X4J camera is large enough. If any larger size is used, the plate- 
holder must be “kitted” down to 3|X4, the standard size of lantern 
slides in America. In using the larger cameras, mark upon the ground 
glass the exact size and location of the lantern-slide plate. Fasten 
the negative in some convenient place where the light may shine 
through it: diffuse daylight is good. Then arrange the camera just 
as in taking any ordinary picture. The board shown in Figure 29 
will be just as useful in making lantern slides as in making photo¬ 
micrographs. At one end of the board fasten a frame which will hold 
an 8X10 negative and also hold kits for smaller negatives (Figs. 29 B 
and C). The long slot in the board will allow the camera to be 
fastened at the proper distance. If buildings, trees, or shadows 
are in the way, tilt the board so as to have a clear sky for a back¬ 
ground. 

Be very careful in focusing; it is best to examine, with a pocket 
lens, the image on the ground glass. In general, use a rather small 
stop, F16 or even F32. If reducing from an average 5X7 negative, 
in good daylight, with an FI6 stop, try 2 or 3 seconds. If enlarging 
from a negative somewhat smaller than a lantern slide, try 8 or 10 


154 


METHODS IN PLANT HISTOLOGY 


seconds. Other things being equal, the best lantern slides are made 
by reduction of larger negatives and the poorest by enlargement 
from smaller negatives. 

The superiority of the larger negative is easily demonstrated. 
With a 5X7 camera, make a negative of some elaborately ornamented 
building, making the building just cover the plate. Then, with a 
3iX4J camera, make a similar negative, so that the building just 
covers the plate. Make lantern slides from both negatives. While 
the building, as it appears on the lantern slide, and on the screen, 
is of the same size in the two cases, the one from the 5X7 negative 
will show much finer detail. The same principle holds true for 
all kinds of plant subjects. The small cameras are easy to carry, 
make good views of interesting bits of scenery, on a tripod, well 
stopped down, will do some fairly good scientific work; but for 
real scientific investigation, take a 5X7. If you are fond of work, 
or can afford to have someone else carry the heavy load, take an 
8X10. 

3 Copying Illustrations.—It is often desirable to get lantern 
slides from photographs, maps, or pictures in books. Here, it is 
necessary to make a negative and then make the lantern slide from the 
negative. In such cases make a 3^X4 negative and print the lantern 
slide by contact. A lantern-slide plate is good for such copying. 
The exposure will depend upon the light, the character of the print, 
and the amount of reduction or enlargement. Other things being 
equal, the exposure will always be longer in case of enlargement than 
in case of reduction. If an average 5X7 photograph is to be copied 
in good diffuse daylight, with an FI6 stop and a lantern-slide plate, 
try 15 seconds. 

For making a negative of a line drawing, about natural size, 
with a Cramer lantern slide for a plate, at F16, dull day with no 
shadows, try 15 seconds. 

For a diagrammatic line drawing, 16X20 inches, F16, good light, 
Cramer lantern slide for a plate, try 10 seconds. 

If one prefers to use film, Eastman Process film is very good for 
copying line work. With the proper exposure and correct develop¬ 
ment it gives excellent results, the lines appearing clean, sharp, 
distinct, with the background perfectly black in the negative. Use 
the developer suggested by the manufacturer. In copying maps 
and line drawings where dead blacks and pure whites are desired, 


PHOTOMICROGRAPHS AND LANTERN SLIDES 


155 


expose fully, but do not overexpose, and develop until the image 
shows plainly on the back of the plate. In such cases, the usual 
developers will not give as good results as a developer modified to 
give dead blacks upon a perfectly transparent background. For 
this purpose, Dr. Land’s formula is the best we have tried. 

Land’s Contrast Developer.—- 


Hydrokinon. 20 g. 

Sodium sulphite (dry). 60 g. 

Sodium carbonate (dry). 140 g. 

Potassium bromide. 12 g. 

Water. 1,000 c.c. 


If kept tightly stoppered, with no air space between the liquid 
and the cork, this developer keeps almost indefinitely. When 
some is taken out for use, the rest should be put into a smaller bottle, 
so that there shall be no air between the liquid and the cork. 

Another formula, developed by Dr. Land to meet the trying 
conditions of the tropics, is also useful for lantern slides. 


Land’s Tropical Developer.— 

Hydrokinon. 

Metol. 

Sodium sulphite (dry). 

Sodium carbonate (dry) 

Potassium bromide. 

Water. 


. 8 g. 

. 3 g. 

. 30 g. 

(60 g. if crystals are used) 

. 30 g. 

(90 g. if crystals are used) 

. 2 g. 

. 1,000 c.c. 


This formula will develop an underexposed plate when the usual 
developers fail. With this developer, the image flashes into sight 
with surprising suddenness, but do npt become startled and remove 
the slide too soon, lest you fail to secure details. 

It is not necessary to furnish the usual formulas for developers 
and fixing solutions, since these are furnished with every box of plates. 
We have found the Cramer plates very satisfactory for all kinds of 
photographic work. The firm will send gratis to anyone who requests 
it Cramer’s manual on Negative-making and Formulas (G. Cramer 
Dry Plate Co., St. Louis, Mo.). 













156 


METHODS IN PLANT HISTOLOGY 


Warm Tones.—A pyro-ammonia developer for warm tones is 
recommended in Harrington’s Photographic Journal for June, 1914: 



Metric Apothecaries’ 

A. Pyro. 

... 31 

g- 

(1 oz.) 

Sodium sulphite crystals.... 

... 62 

g- 

(2 oz.) 

Citric acid. 

... 2.6 g. 

(40 g.) 

Water. 

... 237 

c.c. 

(i pt.) 

B. Ammonia. 

... 31 

g- 

(1 oz.) 

Water... 

... 237 

c.c. 

(J Pt.) 

C. Ammonia bromide. 

... 31 

g. 

(1 oz.) 

Water. 

... 237 

c.c. 

(2 pt.) 

solutions A, B, and C keep well separately, 

but not when 


mixed. When wanted for immediate use, mix A, 1.8 c.c. (30 minims), 
B, 3.7 c.c. (60 minims), and C, 1.8 c.c. (30 minims), and add 30 c.c. 
(1 oz.) of water. 

A correctly exposed plate will develop in about 2\ minutes, and 
the tone should be a warm black. Brown tones are secured by 
increasing the quantity of C, while A and B remain the same. 

Reducing Overexposed Negatives and Lantern Slides.—In case 
of overexposure, the negatives or lantern slides can be saved by 
reducing. The reducing solution should be applied as soon as the 
negative is well fixed in hypo. If a negative which has been washed 
and dried is to be reduced, it should be soaked in water for half an 
hour before using the reducing solution. 

The following is a good solution for most purposes: 


Metric Apothecaries’ 

^ ( Water.. 473 c.c. (16 oz.) 

\ Hyposulphite of soda. 31 g. ( 1 oz.) 

g ( Water. 473 c.c. (16 oz.) 

\ Red prussiate of potassium. 31 g. (1 oz.) 


Solution B must be protected from the light. Cover the bottle 
with black paper and keep it in the dark when not in use. 

Mix only for immediate use 8 parts of A to 1 of B and use in rather 
subdued light. A darkroom is not necessary, but avoid bright 
light. 

When the negative or lantern slide becomes satisfactory, wash 
it in water as thoroughly as if it had just come from the ordinary 
hypo fixing solution. 













PHOTOMICROGRAPHS AND LANTERN SLIDES 


157 


Ordinarily, if it is possible to do so, we believe it is better to make 
a new, correctly exposed negative or lantern slide rather than to 
reduce or intensify an incorrectly exposed one. 

Intensifying Underexposed Negatives and Lantern Slides. —Even 
if a negative or lantern slide has been considerably overexposed, 
it can be reduced quite satisfactorily; if much underexposed, little 
can be done for it; if only slightly underexposed, it may be greatly 


improved by the following solution: 





Metric 

Apothecaries’ 

1 

f Bichloride of mercury. 

2g. 

( 31 gr.) 

A 

Water. 

100 c.c. 

( 4 oz.) 

1 

1 Bromide of potassium. 

2g. 

( 31 gr.) 

B J 

f Sulphite of soda crystals. 

10 g. 

(154 gr.) 

[ Water. 

100 c.c. 

( 4 oz.) 

The solutions keep indefinitely and 

may be 

used three 


times. 

Apply the intensifier after fixing in hypo and washing in water. 
If the negative or slide has been allowed to dry, soak it in water for 
half an hour before intensifying. 

Place the negative or slide in A, rocking the tray as in developing, 
until it becomes gray or even white. Wash in water for 1 minute 
and then transfer to B and leave until the dark color can be seen on 
the back of the negative or slide. Wash in water as thoroughly as 
after fixing in hypo. 

Some use a saturated aqueous solution of the bichloride of mer¬ 
cury, without the bromide of potassium; and, instead of solution B, 
use water to which ammonia has been added—about 1 part ammonia 
to 40 parts water. Excellent sepia tones may be secured in this way. 
Wash well in water. 

After the plate has been thoroughly washed in water, wipe it 
gently with a tuft of cotton. The cotton must, of course, be thor¬ 
oughly wet; it is better to hold the plate under a stream of water 
while wiping. This should always he done before placing a negative or 
slide in the rack to dry, after a washing in water. 

Toning Lantern Slides. —A lantern slide may sometimes be made 
more effective by judicious toning. The hints given here merely 
introduce the student to the possibilities of the subject. 

Light Sepia to Red Tones. —Overexpose up to four or five times 
the length of exposure for a normal slide in black and white; develop 






158 


METHODS IN PLANT HISTOLOGY 


thoroughly; fix and wash as usual; then tone in the following 
solution: 


A. Potassium ferricyanide 

Water. 

B. Copper sulphate. 

Potassium citrate. 

Water. 


Metric 

Apothecaries’ 

6 g. 

( 90 gr.) 

295 c.c. 

( 10 oz.) 

7g. 

( 110 gr.) 

65 g. 

(1,000 gr.) 

295 c.c. 

( 10 oz.) 


(The metric and U.S. measures are practically rather than arithmetically 
equivalent.) 


When needed for use, pour some of A into an equal quantity of 
B, stirring or shaking constantly. Put the slide into the solution in a 
tray and rock just as if developing a plate. The solution is a strong 
reducer. The tone should change from black to warm, then to sepia, 
and may finally become quite red. The time may vary from 1 to 20 
minutes, according to the density of the slide and the tone desired. 
The finished product must not be too dense, for a slide, toned in this 
way, may seem rather weak and yet appear surprisingly strong 
on the screen. 

After toning, wash in water for about 20 minutes. 

Moonlight Tints. —Some excellent formulas, recommended by 


Anderton, will be of service: 

Metric Apothecaries’ 

Ferric ammonia citrate (10 per cent solu¬ 
tion) . .. 15 c.c. (| oz.) 

Potassium ferricyanide (10 per cent solu¬ 
tion) . 15 c.c. (£ oz.) 


Glacial acetic acid (10 per cent solution) 148 c.c. (5 oz.) 

The following gives a more greenish-blue tint: 

Uranium nitrate (10 per cent solu- 


tion in water). 

Ferric ammonia citrate (10 per cent 

3.6 c.c. 

(1 dram) 

solution in water). 

3.6 c.c. 

(1 dram) 

Potassium ferricyanide (10 per cent 

solution in water). 

Nitric acid (10 per cent solution in 

7.2 c.c. 

(2 drams) 

water). 

7.2 c.c. 

(2 drams) 


Both solutions intensify considerably, so that slides to be toned 
should be rather weak. After toning, wash 20 minutes in water. 













PHOTOMICROGRAPHS AND LANTERN SLIDES 


159 


Green Tones.— 


Potassium bichromate (10 

Metric 

per 

Apothecaries’ 

cent solution in water).... 

... 60 

drops 


Potassium ferricyanide (10 

per 



cent solution in water). . . 

. . . 30 

c.c. 

( 1 OZ.) 

Water. 

... 120 

c.c. 

( 4 oz.) 

Cobalt chloride. 

... 3.9 

g- 

(60 gr.) 

Ferric sulphate. 

... 3.9 g. 

(60 gr.) 

Hydrochloric acid. 

... 15 

c.c. 

(h oz.) 

Water. 

... 120 

c.c. 

(4 oz.) 


Bleach in A, wash 10 minutes in water, tone in B, and then 
wash 20 minutes in water. 

Staining Lantern Slides. —Some of the stains used in staining 
microscope slides will give a pleasant tone to lantern slides. Light 
green gives a clear, moonlight effect. Magdala red gives a transparent, 
rosy tint. Sepia and other tones could doubtless be imitated by 
this easy method. 

Clearing Lantern Slides. —Sometimes a slide will seem perfectly 
clear, just as it comes from the fixing bath, especially from an acid 
fixing bath; usually, however, it will be better to transfer the slide 
from the fixing bath to a weak solution of acetic acid—just enough 
acid to give the solution the taste of weak vinegar—and then rock 
for a minute before washing. 

The following clearing fluid may be used in the same way: 


Metric Apothecaries’ 

Alum. 20 g. (1.3 gr.) 

Iron sulphate. 20 g. (1.3 gr.) 

Citric acid. 20 g. (1.3 gr.) 

Water. 500 c.c. (17 oz.) 


Coating Lantern Slides. —After the slide has become thoroughly 
dry, a coat of balsam or shellac will add much to its brilliancy. 
Dilute the Canada balsam with xylol until it becomes almost as 
thin as water; balance the slide on the thumb and first, second, and 
third fingers, holding it as level as possible; pour the balsam over 
it, letting the balsam flow evenly over the whole surface; then tilt 
the slide and pour the balsam back into the bottle. Put the slide in 
the rack to dry. 

Mounting. —Add a suitable mat and a clean lantern-slide cover. 
Remember that the effect of a first-class lantern slide may be impaired 













160 


METHODS IN PLANT HISTOLOGY 


or even ruined by an inartistic mat. Bind the slide and cover together 
with a lantern-slide binding strip. Paste on the label, or, if you 
prefer, paste the label on the mat before binding, so as to have it 
protected by the cover. Lay the slide down so that the positions 
are just as they were in the original, and then paste the “thumb 
mark” in the lower left-hand corner. 

Some efficient and easily available help in making lantern slides 
and in general photography will be found in some booklets published 
by the Eastman Kodak Company, Rochester, New York: “Lantern 
Slides; How to Make and Color Them”; “Elementary Photographic 
Chemistry;” “Color Plates and Filters”; “About Lenses.” 



I 






PART II 


\ 










































SPECIFIC DIRECTIONS 


In the preceding chapters the principles and methods of technic 
have been described in a general way. It is difficult, especially for 
a beginner, to apply general principles to specific cases, and, besides, 
the types which he might select for the preparations might not form 
a symmetrical collection. Consequently, a series of forms has been 
selected which will not merely serve for practice in microscopical 
technic, but will also furnish the student with preparations for a 
fairly satisfactory study of plant structures from the algae up to the 
angiosperms. It is not at all our purpose to discuss general morphol¬ 
ogy, but rather to answer, by means of sketches and specific directions, 
the multitudinous questions which confront the instructor in the 
laboratory. For those who have had a thorough training in general 
morphology the following suggestions will be in some degree super¬ 
fluous. Those who are beginning the study of minute plant structure 
are referred to the standard textbooks for descriptions of the plants 
mentioned here. 

The directions for collecting and growing laboratory material 
constitute an important feature of this part of the book. 

With a few exceptions, the order in which the forms are pre¬ 
sented is that given in Engler’s Syllabus der Pflanzenfamilien. 


163 


CHAPTER XIV 

MYXOMYCETES AND SCHIZOPHYTES 
MYXOMYCETES 

With the exception of a few forms like Fuligo (often found on 
oak stumps and on oak bark in tanyards), the myxomycetes are 

small, and are usually overlooked 
by collectors (Fig. 30). A careful 
examination of rotting logs in 
moist woods will usually reveal 
an abundance of these delicate 
and beautiful organisms. Various 
species may be found in spring, 
summer, and autumn. The plas- 
modia are most abundant just 
after a warm shower. A couple of 
days of dry weather will then 
bring sporangia in abundance. The 
specimens should be pinned to the 
bottom of the box for safe carrying. 
An excellent collecting-box can be 
made from an ordinary paper shoe- 
box. On the bottom of the box 
place a thin piece of soft pine, or a 
piece of the corrugated paper so 
commonly used in packing; or, 
better still, a sheet of cork. At each 
end nail in a piece of pine half an 
inch thick and an inch high. Upon 
these end pieces place a thin piece 
of pine, thus making a second bot¬ 
tom, which, of course, should not be 
fastened. A second pair of ends 
with a third pine bottom nailed to them may rest upon the second 
bottom. The three bottoms will give a considerable surface upon 

164 



gium with some of the rotten wood. The 
dots represent fairly well the size and distribu¬ 
tion of the nuclei at this stage. X70. C, D, 
and E, successive stages, much later than B, 
showing condensation of the wall, origin of 
elaters (D), and mature sporangium (E). 
Preparation stained in safranin, gentian-violet, 
orange. X320. 












MYXOMYCETES AND SCHIZOPHYTES 


165 


which the material may be pinned. For most purposes, the specimens 
are simply allowed to dry, and are then fastened with glue or paste to 
the bottom of a small box. 

Plasmodia and young sporangia may be fixed in chromo-acetic 
acid or Flemming’s fluid. Sections are easily cut in paraffin, and 
should not be more than 5 ju in thickness; for nuclear details, sections 
should not be thicker than 3 /jl. The safranin, gentian-violet, orange 
combination is good for a study of the general development and for 
some cytological features, but iron-alum haematoxylin is better for 
nuclear details. 

Spores of most myxomycetes will germinate as soon as they are 
thoroughly ripe, and, during the first year, germination is more prompt 
than in case of older spores. Fresh spores may germinate in half 
an hour; the time may extend to several hours; spores two or three 
years old may germinate in three or four days, or may not germinate 
at all. We have never succeeded in germinating spores which were 
more than three years old. The longevity is doubtless different 
in different species. In most cases, spores will germinate in water, 
if they will germinate at all. For small cultures, the hanging-drop 
method, described on page 77, may be used. 

Plasmodia may be raised by sowing spores on moist, rotten bark 
or wood and placing the culture under a bell jar, where the moist, 
sultry condition favorable to their growth is easily imitated. Plas¬ 
modia may be got upon the slide by inclining the slide at an angle of 
about 15°, with one end of the slide at the edge of the plasmodium, 
and allowing water to flow very gently down from the upper end of 
the slide to the lower. The proper flow of water could be secured 
by dropping water from a pipette, but a less tedious plan is to arrange 
a siphon so as to secure a similar current. The plasmodium will 
creep up the slide against the current, furnishing an excellent illus¬ 
tration of rheotropism. Enough plasmodium for an illustration may 
be formed in two or three hours. Examined under the microscope, 
the preparation should give an excellent view of the streaming 
movements of protoplasm. 

The following is another method for getting the plasmodia upon 
the slide: Place the slides upon a pane of glass and upon each slide 
place a small piece of plasmodium-bearing wood. Cover with a bell 
jar. Wet blotting paper or a small dish of water included under the 
jar will help to create the warm, sultry atmosphere necessary. The 


166 


METHODS IN PLANT HISTOLOGY 


slides may be covered with the plasmodium in a few hours. Perma¬ 
nent preparations may be made by immersing the slide in chromo- 
acetic acid, then washing and staining without removing the 
plasmodium from the slide. Acid fuchsin is a good stain for 
bringing out the delicate strands of the plasmodium. Iron-alum 
haematoxylin, followed by acid fuchsin or erythrosin, brings out 
both nuclei and cytoplasmic strands. 

Some of the foregoing methods are taken from an article by 
Professor Howard Ayers in the January and February (1898) 
numbers of the Journal of Applied Microscopy. Other methods, 
with directions for various experiments, are given in the same 
article. 


SCHIZOPHYTES (Fission Plants ) 

BACTERIA ( Schizomycetes , Fission Fungi) 

The methods of modern bacteriological technic are so numerous 
and so specialized that we must refer to laboratory manuals for 
instruction in this subject. The method given here will merely 
enable the student to study the form and size of those bacteria 
which are more easily demonstrated. 

Foul water at the outlets of sewers and such places will usually 
afford an abundance of Coccus, Bacillus, Spirillum , and Beggiatoa 
forms. Place a drop of water on a slide, heat it gently until the water 
evaporates, then stain with fuchsin or methyl violet, dehydrate, 
clear in xylol, and mount in balsam. 

The hay infusion is a time-honored method for securing bacteria 
for study. Pour hot water on a handful of hay, and filter the fluid 
through blotting paper. Place the fluid in a glass dish, and cover 
with a piece of glass to keep out the dust. When the fluid begins 
to appear turbid, bacteria will be abundant. The active movements 
are easily observed in a mount from the turbid water. As the 
bacteria pass into the resting condition, they form a scum on the 
surface of the water. Usually, the first to appear is a somewhat 
rod-shaped form, the Bacterium termo of the older texts. Spirillum 
and Coccus forms often appear later. 

Fine preparations may be obtained by inoculating a mouse with 
Anthrax, and then cutting paraffin sections of favorable organs. 
For making mounts of a dangerous form like Anthrax, secure properly 
fixed material from a bacteriologist. Stain in gentian-violet or 


MYXOMYCETES AND SCHIZOPHYTES 


167 


crystal-violet. The following schedule gives good results with 
Anthrax and many other bacteria: 

1. Gentian-violet, 5 minutes. 

2. Rinse in water a few seconds. 

3. Gram’s solution (iodine 1 g., potassium iodide 2 g., water 300 c.c.) 
until the color is almost or quite black; this will generally require 
1 or 2 minutes. 

4. 95 per cent alcohol until the color has nearly disappeared. 

5. Rinse in water and examine. If the bacteria are well stained, a 
counter-stain may be added. 

6. Light green or erythrosin, 5 seconds; or Bismarck brown, 5 or 10 
seconds. 

7. 95 and 100 per cent alcohol, dehydrating as rapidly as possible. Not 
more than 5 or 10 seconds can usually be allowed. 

8. Xylol, 1 to 5 minutes. 

9. Balsam. 

After the rinsing in water of stage 5, the preparation may be 
dehydrated rapidly in 95 per cent and 100 per cent alcohol, and then 
stained for 5 or 10 seconds in orange 
dissolved in clove oil. From the 
clove oil, transfer to xylol and mount 
in balsam (Fig. 31). 

With sections not more than 5 /jl 
thick, excellent results can be ob¬ 
tained by staining in iron-alum 
haematoxylin. 

The following rapid method gives 
fairly good results: 

1. Place on a clean cover a drop of 
water containing the bacteria and 
dry completely in a flame or on a 
hot plate. 

2. Stain 2 to 5 minutes in gentian- 
violet or methyl violet. 

3. Rinse quickly in water. 

4. Dip into 95 per cent alcohol to 
reduce the stain. 

5. Remove most of the alcohol by 
touching a corner of the cover with 
filter paper and then dry completely by passing through a flame. 

6. Mount in balsam. 



Fig. 31 .—Bacillus anthracis, from a 
paraffin section cut from the liver of a 
mouse and stained in crystal-violet; w, 
white blood corpuscle; r, red blood cor¬ 
puscle. X 580. 




168 


METHODS IN PLANT HISTOLOGY 


Leptothrix may often be obtained by scraping the inside of the 
cheek. Beggiatoa, one of the sulphur bacteria, with oscillating 
movements like Oscillatoria, is often found in foul water. Its presence 
may be indicated by whitish patches on the bottom. 

The Bacteria are the only plants in which a nucleus has not been 
conclusively demonstrated, and some claim that a nucleus is present 
even in Bacteria. In determining the presence or absence of a 
nucleus in Bacteria, the crude method, just given, would be of no 
value, and even the most critical methods of the bacteriologist, who 
mounts the organisms whole, would be entitled to only scant considera¬ 
tion. The presence or absence of a nucleus will have to be determined 
by a study of thin, well-stained sections of perfectly fixed material. 

CYANOPHYCEAE. BLUE-GREEN ALGAE (,Schizophyceae Fission Algae ) 

The blue-green algae include unicellular, colonial, and filamentous 
forms. They occur everywhere in damp or wet places. On the 
vertical faces of rocks where there is a constant dripping of water, 
brilliant blue-green forms are abundant. In the Yellowstone National 
Park the brilliant coloring of the rocks is due in large measure to 
members of this group. Many forms occur as brownish or greenish 
gelatinous layers on damp ground or upon rocks, or even upon 
damp wooden structures in greenhouses. Other forms float freely 
in water or on the surface of the water. 

Oscillatoria.—For most purposes it is best to study Oscillatoria 
in the living condition. It is readily found in watering-troughs, in 
stagnant water, on damp earth, and in other habitats. The com¬ 
monest forms have a deep blue-green or brownish color. It is very 
easy to keep Oscillatoria all the year in the laboratory. Simply 
put a little of a desirable form into a gallon glass jar half filled with 
water. By adding water occasionally to compensate for evaporation, 
the culture should keep indefinitely. In a jar with a tightly fitting 
cover we have kept such a culture for years without renewing the 
water. 

For the purposes of identification and herbarium specimens the 
material may simply be placed on a slip of mica and allowed to dry. 
When wanted for use, add a drop of water and a cover, and the mount 
is ready for examination. After the examination has been made 
remove the cover, allow the preparation to dry, and then return it 
to the herbarium. 


MYXOMYCETES AND SCHIZOPHYTES 


169 


Good mounts may be made by the Venetian turpentine method. 
Species of medium size are more satisfactory for a study of the 
nucleus than the very large species. Fix in a chromo-acetic-osmic 
solution (1 g. chromic acid, 3 c.c. acetic acid, and 1 c.c. 1 per cent 
osmic acid). Stain in iron-alum haematoxylin, and follow the 
Venetian turpentine method. While the nuclei are easily seen in 
such preparations, still better views can be secured from sections of 
paraffin material fixed in the same solution or in Flemming’s weaker 
solution. The section should be about 3 p thick. After staining with 
haematoxylin, stain lightly with orange dissolved in clove oil. In 
paraffin sections the scattered condition of the material as it appears 
in thin sections is very annoying. As soon as the material is thor¬ 
oughly washed in water, arrange it so that the filaments will all have 
the same general direction. This will enable you to get longitudinal 



Fig. 32.— Oscillatoria: photomicrograph from a paraffin section 3 n in thickness and stained 
in iron-alum haematoxylin. X373. 

and transverse sections. As you begin with the alcohols, use a Petri 
dish and lay a slide over the material, and keep it there until you 
imbed in paraffin. This will keep the filaments from spreading out 
too much, and you will be able to get as much on one slide as you 
would be likely to get on a dozen slides without such precaution. 

Oscillatoria , as it appears in section, is shown in Figure 32. 

Tolypothrix.—This form occurs as small tufts, either floating 
in stagnant water or attached to plants and stones. Some species 
grow upon damp ground. It furnishes an excellent example of false 
branching (Fig. 33). Like all small filamentous algae, it may be 
dried on mica for herbarium purposes. Venetian turpentine mounts 
and paraffin sections are prepared as in Oscillatoria. Tolypothrix is 
even better than Oscillatoria for a study of the nucleus. 

Scytonema is a similar form which is fairly common. It is often 
found as a feltlike covering on wet rocks. 







170 


METHODS IN PLANT HISTOLOGY 


In staining forms like Tolypothrix and Scytonema, which have a 
thick sheath, take care not to obscure the cell contents by staining 
the sheath too deeply. If the sheath is not stained at all, you may 
not be able to see the nature of the false branching. Iron-alum 


haematoxylin, with orange in clove oil for the sheath, is good for sec¬ 
tions. Magdala red, with light green for the 
sheath, is good for Venetian turpentine mounts. 

Nostoc.— Nostoc is a cosmopolitan form. It 
occurs on damp earth or floating freely in water. 
In a fruit can or a battery jar, Nostoc is easily 
kept year after year in the laboratory. Young 
specimens are generally in the form of gelatinous 
nodules, but in older specimens the form may 
be quite various. It is very easy to make 
sections, since the gelatinous matrix cuts well 
and holds the filaments together. Chromo- 
acetic acid is a good fixing agent. Stains which 
stain the gelatinous matrix make the preparations 
look untidy, but they show that each filament 
of the nodule has its own gelatinous sheath. 
Small nodules may be stained in bulk and be 
got into Venetian turpentine. Crushed under 
the cover, they make instructive preparations. 

Rivularia.—-This form is readily found on the 
underside of the leaves of water-lilies (. Nuphar , 
Nymphaea, etc.), but is also abundant on sub¬ 
merged leaves and stems of other plants. It occurs 
in the form of translucent, gelatinous nodules of 
various sizes. Chromo-acetic acid gives beautiful preparations, but 
good results can also be secured from formalin or picric-acid material. 

The most instructive preparations for morphological study can 
be obtained by the Venetian turpentine method. Stain in iron- 
haematoxylin and very lightly in erythrosin, the latter stain being 
used merely to outline the sheath. When ready for mounting, 
crush a small nodule under a cover-glass. The paraffin method is 
easily applied, since the gelatinous matrix keeps the filaments in 
place. Any form of similar habit may be prepared in the same way. 

Gloeotrichia — Gloeotrichia (Fig. 34), in its later stages, is a free- 
floating form. In earlier stages it is attached to various submersed 


Fig. 33.— Toly-pothrix f 
showing “false branch¬ 
ing”: h, heterocyst; c, con¬ 
cave cell; 6, end of false 
branch with beginning of 
new sheath. X620. 


















MYXOMYCETES AND SCHIZOPHYTES 


171 


aquatic plants. The nodules, when young, are firm like Nostoc , 
but as they grow older and larger they become hollow and soft. 
The older forms become so much dissociated that they lose their 
characteristic form and merely make the fixing fluid look turbid. 
Allow a drop of such material to spread out and dry upon a slide 
which has been slightly 
smeared with albumen 
fixative. Leave the 
slide in 95 per cent 
alcohol 2 or 3 minutes 
to coagulate the albu¬ 
men fixative, and then 
stain in safranin. If 
the background ap¬ 
pears untidy, stain for 
24 hours, or longer; 
you can then extract 
the stain from the 
background, and still 
leave the long spore 
and some of the other 
features of the fila¬ 
ment well stained. A 
touch of cyanin will 
bring out the sheath. 

Cyanin and erythro- 
sin is a good combina¬ 
tion if the material 
is clean. The firmer nodules may be treated like Nostoc or Rivularia. 

Wasserbliithe.—Many genera of the Cyanophyceae occur as 
scums, often iridescent, on the surface of stagnant or quiet water. 
Some of the commonest forms are Coelosphaerium and Anabaena 
(Fig. 35). Some of the Chlorophyceae also occur as Wasserbliithe. 
Where the material is very abundant, it may be collected by simply 
skimming it off with a wide-mouthed bottle, but where it is rather 
scarce, it is better to filter the water through bolting silk and finally 
rinse the algae off into a bottle, adding enough formalin to the water 
in the bottle to make a 5 per cent solution. The material may be 
kept here indefinitely, but after a few hours it is ready for use. If 



Fig. 34.— Gloeotrichia: photomicrograph from a preparation 
stained in cyanin and erythrosin: negative by Dr. W. J. G. Land. 



172 


METHODS IN PLANT HISTOLOGY 


the forms are small, like Anabaena, smear a slide lightly with Mayer’s 
albumen fixative, as if for paraffin sections, add a drop of the material 
and allow it to dry over night or for 24 hours; then immerse the slide 
in strong alcohol for a few minutes, and then proceed with the 
staining. Cyanin and erythrosin form a good combination for differ¬ 
entiating the granules. Delafield’s haematoxylin, used alone, stains 
some granules purple and others red. Iron-alum haematoxylin 
is excellent for heterocysts. With patience, these Wasserbliithe 
forms may be stained in iron-haematoxylin and brought into Venetian 
turpentine, from which they will yield much better preparations 

than can be secured by the drying- 
down method. 

Sometimes Anabaena, mixed 
with Gloeothece or Gloeocapsa, occur 
floating in gelatinous masses which 
hold together fairly well, so that 
it is easy to fix in the chromo- 
acetic-osmic solution recommended 
for Oscillatoria, stain in iron-alum 
haematoxylin, and follow the Ve¬ 
netian turpentine method. 

With such material we have 
tried a more expeditious method 
with excellent results. After staining in haematoxylin, we have 
used a series of alcohols, 2J, 5, 10, 15, 20, 30, 40, 50, 70, 85, 95, 
and 100 per cent, allowing only 3 or 4 hours for the entire series. 
Then use mixtures of clove oil and absolute alcohol, beginning 
with 1 part clove oil to 4 parts alcohol, followed by equal parts 
clove oil and alcohol, then 3 parts clove oil to 1 of alcohol. At 
this point, stain in orange dissolved in clove oil. Drain off the 
stain and transfer to pure clove oil. Then place the material in 
thin balsam, about 1 part of the balsam used for mounting to 3 parts 
of xylol. Here the material may be kept indefinitely. Mounts 
may be made in balsam from this stock. Figure 35 was drawn from 
material prepared in this way. 



Fig. 35.— Anabaena: A, hormogonium 
showing well-defined nuclei; B, older filament 
showing a spore and a heterocyst. 


CHAPTER XV 

CHLOROPHYCEAE. GREEN ALGAE 

For experiments in most phases of botanical microtechnic, no 
group of plants affords better material than the green algae, since the 
killing, fixing, and staining can be watched directly; the effect of 
the change from one solution to another can be observed; and even 
the behavior during infiltration with paraffin can be determined with 
considerable accuracy. 

Since the Chlorophyceae furnish our best illustrations of the 
evolution of the plant body, the origin and development of sex, and 
also the beginning of alternation of generations, they occupy a 
prominent place in any well-planned course in the morphology of 
plants; and, if they were better known, the ease with which the 
reactions of the individual cell may be observed would make them 
valuable to the physiologist. 

They are found in both fresh and salt water, but are most abun¬ 
dant in fresh water. The ponds, ditches, and rivers of any locality 
will yield an abundance and variety both of the unicellular and the 
multicellular members of this group. Most of the forms are inde¬ 
pendent, but there are epiphytic, endophytic, and saprophytic 
species. The larger forms and those which grow in tufts or mats are 
readily recognized in the field. Many of the smaller forms are 
attached to other water plants. Drain the water plants and then 
squeeze them over a bottle. The sediment is likely to contain a 
variety of unicellular and other small algae. 

Many of the genera are easily kept in the laboratory. It is not 
necessary to have very large aquaria. Ordinary glass battery jars 
holding about a gallon are good for most forms. Jars holding 2 gal¬ 
lons will be as good or better. For some cultures which are to 
be kept for a long time, like Scenedesmus, small glass jars, or dishes, 
with ground-glass tops are desirable. For a limited amount of 
material, quart or 2-quart fruit cans are very efficient. Put about 
an inch of pond dirt in some, clean sand in others, and in still others 
use a gravel bottom. Many forms grow well without any soil or 
sand in the bottom of the jar. 


173 


174 


METHODS IN PLANT HISTOLOGY 


When possible, use the water in which the algae were growing, 
since very few take kindly to a sudden change of water. If the 
material has been brought to the laboratory in a very small quantity 
of water, fill the jar about two-thirds full with tap water. Let the 
water run for 2 or 3 minutes before you fill the jar, since the water 
standing in the pipes is injurious, or even fatal, to most algae. Add 
water occasionally, only a little at a time, to compensate for evapora¬ 
tion. If the water has evaporated until the jar is about one-third 
full and you fill it nearly to the top with tap water, you are likely 
to kill some of the most desirable forms. 

It is a mistake to put too much material into a jar. A wad of 
Spirogyra half as large as one’s finger is as much as should be put 
into a gallon jar. As it grows to ten or twenty times that amount, 
it is not necessary to keep throwing it out, since it will gradually 
accommodate itself to conditions. However, do not let the jar 
become choked with the material. 

Cultures may be started even in the winter. Bring in some mud 
over which algae were growing the previous summer or autumn; 
put it into a jar and fill it two-thirds full of tap water. Also bring 
in sticks, leaves, and stones from good alga localities and put them 
into jars of tap water. Cultures may be started either by taking 
mud and sticks from under the ice or by taking them from places 
which have entirely dried up during the summer or autumn. A 
few such jars will be likely to yield a variety of material. 

If you have a good jar of Oedogonium, or some other desirable 
form, do not throw it out if the alga should disappear. Remember 
that temporary disappearances occur in nature. Allow the culture to 
become dry and then set it aside where it will be protected from dust. 
After a few months, pour on tap water and it is very likely that you 
will soon have a good jar of Oedogonium. Many algae behave simi¬ 
larly; some, like Volvox, appear for a short time and then disappear for 
a long time; some, like Cladophora, may last the whole year and grow 
so luxuriantly that the excess material must be removed; and some, 
like Ulothrix, we have not been able to cultivate at all in the laboratory. 

Some very useful hints on collecting and growing fresh-water 
algae for class work will be found in an article by Dr. J. A. Nieuwland 
in the Midland Naturalist , 1:85, 1909. 

Professor Klebs has shown that various phases in the life-histories 
of many algae and fungi may be produced at will. By utilizing his 


CHLOROPHYCEAE 


175 


results, the fruiting condition may be induced in many of the common 
laboratory types. Knop’s solution will be needed in most cases. 
A stock solution which can be diluted as required may be made as 


follows: 

Potassium nitrate, KN0 3 . 1 g. 

Magnesium sulphate, MgS0 4 . 1 g. 

Calcium nitrate, Ca(N0 3 ) 2 . 3 g. 

Potassium phosphate, K 2 HP0 4 . 1 g. 


Dissolve the first, second, and fourth ingredients in 1 liter of 
distilled water, and then add the calcium nitrate. A precipitate of 
calcium phosphate will be formed. For practical purposes this may 
be called a 0.6 per cent solution. Whenever a dilute solution is 
made from the stock solution, the bottle must be shaken thoroughly 
in order that a proper amount of the precipitate may be included in 
the diluted solution. To make a 0.1 per cent solution, add 5 liters of 
distilled water to 1 liter of the stock solution; for a 0.3 per cent solu¬ 
tion, add 1 liter of distilled water to 1 liter of the stock solution, etc. 

The addition of a liter of a 0.2 per cent solution to 4 or 5 liters 
of water will often produce a more thrifty growth. Directions for 
inducing reproductive phases are given in connection with the various 
types. With a good supply of glass jars, plenty of Knop’s solution, 
a reasonable control over temperature, and the teacher’s usual 
amount of patience, most laboratory types can be studied in the 
living condition at all seasons of the year. 

Collecting algae need not be so laborious as most botanists make 
it. Forms like Spirogyra, Zygnema, Cladophora, Vaucheria , and 
Hydrodidyon may be rolled up in wet newspaper and carried in a 
botany can. They suffer less from lack of water than from lack of 
air. Large quantities of material can be brought in and transferred 
to water after reaching the laboratory. Even after 24 hours in the 
wet paper, such forms seem to suffer no damage. 

Permanent preparations are needed to show details which are 
not so evident in the fresh material. The unicellular and filamentous 
members, together with such forms as Volvox, are best prepared by 
the Venetian turpentine method. The structure is so much more 
complicated than in the Cyanophyceae that it demands far more care 
and skill to make good preparations. 

In many, probably in most of the green algae, nuclear and cell 
division takes place at night. This is definitely known to be the case 






176 


METHODS IN PLANT HISTOLOGY 


in Spirogyra , Zygnema, Closterium, Ulothrix, and others. Mitosis 
is most abundant about midnight, or an hour before midnight, and 
continues up to three or four o’clock in the morning. The most 
extensive work on the time of day at which nuclear division occurs 
is a paper by G. Karsten, “Ueber embryonales Wachstum und seine 
Tagesperiode, Zeitschrift fiir Botanik , 7:1-34, 1915. Although the 
paper is in German, the numerous tables can be understood by 
those who are unfamiliar with the language. The paper contains a 
bibliography of the subject. 

Chromo-acetic acid, with or without osmic acid, is a good killing 
and fixing agent for the entire group. We prefer the following for¬ 
mula : Chromic acid, 1 g.; glacial acetic acid, 3 c.c; 1 per cent osmic 
acid, 1 c.c. . 

With any fixing agent, it is worth while to place a few filaments 
in the mixture and watch the effect under the microscope. If 
plasmolysis occurs with the chromo-acetic mixture, weaken the 
chromic or strengthen the acetic until the suitable proportions are 
determined. In the previous edition, the usual method was to 
weaken the chromic acid. While this avoided any shrinking of the 
cell contents, the fixing was not very thorough, and material often 
suffered during staining or other subsequent processes. An extensive 
series of experiments, especially with coenocytic forms which are 
notoriously difficult to prepare, proved that it is better to keep the 
chromic acid up to 1 per cent and strengthen the acetic acid, if neces¬ 
sary. The function of the osmic acid is to make the killing almost 
instantaneous. This high percentage of acetic acid, excellent for 
algae and fungi and many other forms, is not so good for many tissues 
of higher plants, because the proportion of acetic acid is too great. 
About 24 hours in any of the chromic series and a 24 hours’ washing 
in water will be sufficient for members of this group. Only a few 
of the most commonly studied will be mentioned. 

With Marine Forms use sea-water in making up the fixing agents 
and in washing, but use fresh water in making up alcohols and for the 
10 per cent glycerin. 

Volvox.— Volvox is found in ponds and ditches, and even in 
shallow puddles. The most favorable place to look for it is in the 
deeper ponds, lagoons, and ditches which receive an abundance of 
rain water. It has been claimed that where you find Lemna, you 
are likely to find Volvox; and it is true that such water is favorable, 


CHLOROPHYCEAE 


177 


but the shading is unfavorable. Look where you find Sphagnum, 
Vaucheria, Alisma, Equisetum fluviatile, Utricularia, Typha, and 
Char a. Dr. Nieuwland reports that Pandorina, Eudorina and 
Gonium are commonly found in summer as constituents of the green 
scum on wallows in fields where pigs are kept. The flagellate, 
Euglena, is often associated with these forms. If you have a culture 
in the laboratory, do not throw it out when the culture disappears, 
because new coenobia are likely to develop from the oospores. 

For collecting, use “bolting cloth” or “bolting silk” of the finest 
mesh available. With a piece of thin cloth about 15 cm. square, 
laid over an ordinary coffee strainer, you can pour through about 
4 liters of water in a minute. In this way you will secure all the 
Volvox in about a barrel of water in half an hour. Eudorina may be 
collected in the same way. Smaller members of the Volvox family 
like Pandorina, Gonium, and Chlamydomonas are too small to be held 
by the cloth; but if material is very abundant, the water goes through 
faster than the organisms and you will soon have many times as 
much material in a bottle as you could get by dipping. Many 
small organisms are effectively collected in this way, even when they 
are so small that most of them pass through the cloth. 

Material of Volvox and all the Volvocaceae may be fixed in the 
corrosive sublimate-acetic mixture, used hot—85° C. If material is 
to be stained and mounted whole, use the aqueous mixture; if it is 
to be imbedded and cut, use the alcoholic. For mounting whole, 
stain in iron-alum haematoxylin, or in Magdala red and anilin blue, 
following the Venetian turpentine method. A few bits of broken 
cover-glass, placed among the colonies, will prevent crushing. 

Many fixing agents cause the protoplasmic connections between 
the cells to be withdrawn. Figure 36 A, showing the protoplasmic 
connections in nearly the living condition, was drawn from material 
fixed in the hot aqueous corrosive sublimate-acetic acid solution; 
B, fixed in a 1 per cent chromo-acetic solution, does not show the 
connections; C to E show details drawn from sections. 

For paraffin sections, the material, preferably in sufficient 
abundance to make a layer half an inch deep in the bottom of a bottle 
as large as one’s finger, is infiltrated with paraffin in the usual way. 
In imbedding, simply pour the contents of the bottle out so as to 
form a thin layer on a piece of glass. If material is so abundant 
that you can afford to lose some of it, simply cool the paraffin in 


178 


METHODS IN PLANT HISTOLOGY 


the bottle or dish. If the paraffin sticks, as it probably will, break 
the dish and cut the paraffin into suitable blocks. Or, the paraffin 
may be poured into the usual imbedding dish, keeping it just warm 
enough to let the organisms settle; then cool and cut into blocks. 
Safranin, gentian-violet, orange, are very good for olden stages, 
especially for pyrenoids and starch. Iron-alum haematoxylin is 
better for nuclei and the younger stages of oogonia, antheridia, and 
new colonies. 



Fig. 36.— Volvox: surface view of several cells, fixed in the hot aqueous corrosive sublimate- 
acetic acid mixture, stained in iron-alum haematoxylin, and mounted in Venetian turpentine; B, 
fixed in chromo-acetic acid, but otherwise treated like A; C-G fixed in 1 per cent chromo-acetic 
acid imbedded in paraffin and cut 5 ju; C, E, and F stained in iron-alum haematoxylin; D, and G 
stained in safranin gentian-violet orange; C, new colony showing pyrenoids, p, and nucleus, n; 
D, a nearly mature antheridium; E, young egg; F, egg before fertilization; G, egg after fertilization, 
showing oil globules, o; pyrenoids, p; starch cut off from pyrenoid, s. All X780. 

Figure 37 shows that even such delicate forms as Volvox can be 
imbedded in paraffin without shrinking. 

The Power’s Methods.—-Professor J. H. Power’s mounts of 
Volvox and other members of the Volvocaceae have been the delight 
and despair of both botanists and zoologists for many years. They 
have never been surpassed and, probably, never equaled. Professor 
Powers has kindly given me an outline of his methods; but, as in 
other phases of technic, judgment, skill, and patience must be fur¬ 
nished by the student. 




CHLOROPHYCEAE 


179 


For fixing, Professor Powers uses the aqueous potassium iodide 
solution used in testing for starch. This solution may be weaker 
than the usual formula so that it has a light-brown color. From 
10 to 24 hours is sufficient for fixing, but material may be left here 
for several days. Wash thoroughly in tap water which has stood 
long enough to give off all of its excess of air; otherwise bubbles 
will form on the colonies, causing them to float and hindering 
subsequent processes. Change after change of water should be 
made rapidly, using large 
amounts of water and 
decanting just as soon as 
the colonies have settled. 

From 1 to 3 hours’ wash¬ 
ing should be sufficient to 
remove the brown color 
of the iodine. 

Stain in Mayer’s Car- 
malum. 1 Use a pure car- 
minic acid in making the 
stain, 1 g. carminic acid, 

10 g. alum, and 2Q0 c.c. 
distilled water. Dissolve 
with heat, filter, and add 
a crystal of thymol to 
keep out fungi. After 
staining, follow the Vene¬ 
tian turpentine method material fixed in chromo-acetic acid and stained in Dela- 
^ } field’s haematoxylin; from a preparation and negative by 

taking care to wash the Dr. w. j. g. Land, 
glycerin out completely. 

The 10 per cent turpentine should not be allowed to concentrate too 
rapidly. 

Material fixed in weak osmic acid is even better for protoplasmic 
connections and cilia. About 4 or 5 drops in 50 c.c. of distilled water 
is sufficient. From 6 to 24 hours is long enough for fixing. 

After either of these fixing agents, following the washing in water, 
material may be preserved in a nearly saturated solution of alum; 
or in a dilute aqueous carmalum, with a crystal of thymol to prevent 
mold. Staining for 3 weeks in a weak carmalum stains the cells 

1 Lee Vade-Mecum (8th ed. p. 137). 



Fig. 37.— Volvox: photomicrograph of a section from 


180 


METHODS IN PLANT HISTOLOGY 


but not the matrix. About 3 months will be necessary to stain 
the matrix enough to make it a background for the cells. 

An aqueous solution of nigrosin gives an effect like that of iron- 
alum haematoxylin. Rose benzol and Lee’s pyrogallic acid method 
were also useful. 

Diatoms.—Living diatoms are often found clinging in great 
numbers to filamentous algae, or forming gelatinous masses on vari¬ 
ous submerged plants. Cladophora is frequently covered with 
Cocconeis, an elliptically shaped diatom; Vaucheria is often covered 
with small forms. Other algae will pay for examination, especially 
if they look brown. If stones in the water have a brown, slippery 
coating, you can be sure of diatoms. Sometimes the brown coating 
on sticks and stones is so abundant that it streams out with the 
current. If rushes and stems of water plants have a brown, gelati¬ 
nous coating, you are likely to find millions of specimens of the same 
diatom. The surface mud of a pond, ditch, or lagoon will always 
yield some diatoms. They can be made to come out from the 
mud by putting a black paper around the jar and letting direct 
sunlight fall upon the surface of the water. The diatoms, in a 
day or even less, will come to the top in a scum which can be easily 
secured. 

Since diatoms form an important part of the food of molluscs, 
tunicates, and fishes, the alimentary tracts of these animals often 
yield deep-water forms which are not easily secured in any other way. 

Fresh-water diatoms appear in greatest abundance in spring, 
are comparatively scarce in summer, and reappear in autumn, 
though not so abundantly as in the spring. 

Marine forms can be secured by scraping barnacles, oyster shells, 
and other shells. The big Strombus shell from the West Indies, 
which we use to keep the door open, will yield a good collection if 
you get it before it is cleaned. 

The silicious shells of diatoms are among the most beautiful 
objects which could be examined with the microscope (Fig. 38). 
To obtain perfectly clean mounts requires considerable time and 
patience, but when the material is once cleaned, preparations may 
be made at any time with very little trouble. Diatom enthusiasts 
h^ve devised numerous methods for cleaning them, and separating 
the various forms from one another, but we shall give here only a few 
simple, practical methods. 


CHLOROPHYCEAE 


181 


Material for mounts of frustules of living forms 
may be obtained by skimming off the brownish scum found on ponds, by 
squeezing out water weeds, by scraping sticks and stones which are covered 
at high water, or from the mud of filter beds and pumping-works, or in 
other places. The material is put in a dish of water, and after it has settled 
the water is decanted. This is repeated until the water will clear in about 
half an hour. The sediment is then treated with an equal bulk of sulphuric 
acid, after which bichromate of potash is added until all action ceases. 
After a couple of hours the acid is washed out. To separate the diatoms, 
place the sediment in a glass dish with water, and when the water becomes 
clear give the dish a slight rotary motion. This will bring the diatoms to the 
top, when they may be removed with a pipette and placed in alcohol. To 
mount, place a number in distilled water, evaporate a few drops of the 
mixture on a cover-glass, which is then mounted on a slide in balsam. 1 

It is better to use a very slight smear of Mayer’s albumen fixative 
to prevent the diatoms from floating to the edge of the cover. 

Many scouring soaps and silver polishes contain large quantities 
of fossil diatoms, and the diatomaceous earths are particularly rich. 
Diatomaceous earths from Cherryfield, Maine, and from Beddington, 
in the same region, are the richest we have seen. The deposits at 
Richmond, Virginia, have long been famous. In some of our western 
states there are deposits 300 feet thick, with 80 per cent of silica, 
the silica being the valves of diatoms. 

Break up a small lump of such material and boil it in hydrochloric 
acid. An evaporating dish or a test-tube is convenient for this 
purpose. Let the diatoms settle, pour off the acid, and then wash 
in water. As soon as the diatoms settle, the water should be poured 
off. The washing should be continued until the hydrochloric acid 
has been removed. When the washing is complete, pour on a little 
absolute alcohol, and after a few minutes pour off the alcohol and 
add equal parts of turpentine and carbolic acid. The material will 
keep indefinitely in this condition, and may be mounted in balsam 
at any time. In making a mount, put a little of the material on a 
slide and allow it to become dry, or nearly dry, and then add the 
balsam and cover. If the balsam should be added too soon, the 
diatoms are likely to move to the edge of the cover. 

We have had excellent results with the following method: After 
washing in water, keep the diatoms in 5 per cent formalin. To 

1 From a review of Dr. Wood’s paper on “Diatoms,” Journal of Applied Microscopy, 
March, 1899. 


182 


METHODS IN PLANT HISTOLOGY 



make a mount, smear the slide very slightly with Mayer’s albumen 
fixative, add a little of the material, and heat just enough to coagulate 
the albumen. When perfectly dry, add a drop of balsam and a cover. 
Or, after coagulating the fixative, dip in absolute alcohol and then in 
xylol before mounting in balsam. Without the alcohol and xylol, 


some air is sure to be caught and it may accentuate some markings; 
but, in general, we prefer to use the alcohol and xylol. 

To show the cell contents, diatoms must be fixed and stained. 
If they are clinging to filamentous algae, the algae with the diatoms 
attached should be put into chromo-acetic acid (24 hours) and then 
washed in water for 24 hours. Stain in iron-haematoxylin and 


Fig. 38. —Diatoms: diatomaceous earth from Cherryfield, Maine, a Pleistocene deposit, 
showing the great variety of forms usually found in such material; photomicrograph by Miss 
Ethel Thomas from a preparation by Rev. E. L. Little. X400. 




CHLOROPHYCEAE 


183 


proceed by the Venetian turpentine method. When ready for 
mounting, the diatoms can be scraped off from the algae or other 
substratum. Other stains may be used. 

When the material is in gelatinous masses it may be fixed in 
chromo-acetic acid, with or without a little osmic acid, and imbedded 
in paraffin. There will, of course, be some difficulty in cutting, 
but the knife often breaks the frustules very cleanly, so that good 
sections may be secured. It might be worth while to try a weak 
solution of hydrofluoric acid to dissolve the silicious shells. 

Desmids.—The desmids are unicellular, free-floating or sus¬ 
pended algae. They are not found in salt water and are more 
abundant in soft water than in hard. Deep pools, quiet ponds, and 
quiet margins of small lakes are good collecting-grounds. Collections 
of other fresh-water algae often contain some desmids. It frequently 
happens that a single desirable desmid appears during examination 
of field collections. In such a case, remove it with a fine pipette, 
and get it into a drop of water on a clean slide, invert it over a bottle 
of 1 per cent osmic acid for 2 minutes, leave the slide exposed to the 
air until almost all the water has evaporated, and then add a drop 
of 10 per cent glycerin. In a few hours (6 to 24) put on a cover and 
seal. It requires more time, care, and patience than it is worth to 
attempt staining in such a case. 

Sometimes desmids occur in great abundance. They may then 
be treated like the filamentous algae, except that more care must 
be taken not to lose them when changing fluids. Four or five drops of 
1 per cent osmic acid to 50 c.c. of water fixes well, and material from 
this solution may be placed directly into 10 per cent glycerin and 
mounted by the Venetian turpentine method. It looks almost as 
if stained in iron-alum haematoxylin. The iodine solution used in 
testing for starch gives good results and may be followed by any 
stains. The larger desmids stain beautifully in iron-alum haema¬ 
toxylin. 

The Venetian turpentine method, with Magdala red and anilin 
blue, will give beautiful preparations. A deep stain with Magdala 
red and a rather light stain with anilin blue is better for the pyrenoids 
and nucleus, while a light stain in the red and a deep stain in 
blue is better for the chromatophores. When the material is suffi¬ 
ciently abundant, paraffin sections may be made as directed for 
Volvox. 


184 


METHODS IN PLANT HISTOLOGY 


Lutman has found that Closterium divides at night. If mitotic 
figures are wanted they are likely to be obtained if the material is 
fixed about midnight. 

Zygnema. —Zygnema is one of the commonest algae of the ponds, 
swamps, and ditches. The mats are very slippery to the touch. 
In the field it resembles Spirogyra, but is distinguished by the two 
characteristic chromatophores which are readily seen with a good 
pocket lens. Sometimes conjugation can be induced by bringing 
the material into the laboratory and placing it in open jars with 
plenty of water and not too much light. 

The special chromo-acetic-osmic-acid solution is good for fixing. 
Stain in iron-alum haematoxylin and also in Magdala red and anilin 
blue. Follow the Venetian turpentine method, and in mounting 
put material from both stains on each slide. 

Textbooks describe “ stellate chromatophores ” in Zygnema. 
A good preparation should show that the chromatophores have 
an even outline, with no trace of a stellate form. The boundary 
between the chromatophore and the protoplasm—often of a stellate 
form—in which the chromatophore is imbedded, should be seen in 
well-fixed material with either of the foregoing stains. The large 
starch grains, extending from the pyrenoid almost to the border of 
the chromatophore, are better differentiated b^ the Magdala red 
and anilin blue; the nucleus and pyrenoid are better stained by the 
iron-alum haematoxylin. Careful staining should bring out the 
features shown in Figure 39. The chromatophores do not stain as 
readily as those of Spirogyra , and consequently it is necessary to use 
stronger stains or more prolonged periods. Use the Venetian turpen¬ 
tine method. 

For a detailed study, imbed in paraffin and cut thin sections. 
After washing in water, arrange the filaments so that most of them 
will have the same general direction; then, in running up through 
the alcohols, keep the filaments from spreading too much by placing 
a slide on the material. After imbedding, the material can be cut 
into blocks about a centimeter square. If sections thinner than 5 p 
are wanted, cut out smaller paraffin blocks. 

Spirogyra.—Probably no alga has been more studied by pupils, 
teachers, and investigators than Spirogyra. Nearly all of the 
numerous species belong to the low, quiet waters of ponds and 
ditches, where they often form large, flocculent green mats nearly 


CHLOROPHYCEAE 


185 


covering the surface of the water. A few species occur in running 
water. The mats are very slippery to the touch—a character which 
assists in recognizing the genus in the field. In the larger species 
the characteristic spiral chromatophores can be seen with a good 
pocket lens, thus completing the identification, as far as the genus is 
concerned. Mats in which zygospores have been formed are likely 
to show a pale, or even a brownish, color, due to the brownish walls 



Fig. 39. — Zygnema: fixed in the special chromo-acetic-osmic solution: A and B stained in 
Magdala red and anilin blue; C and D stained in iron-alum haematoxylin. All show the nuclei, 
the chromatophores differentiated from the cytoplasm, and the large starch grains arranged radially 
about the pyrenoids. In C, cell division has just taken place and the pyrenoids and chromatophore 
in each new cell are dividing. In D, a young zygospore, the two nuclei have not yet fused. X790. 


of the zygospores. This color, however, is not always, or even 
usually, due to zygospores, but is more often due to the death and 
degeneration of the plants. Mats in early stages of conjugation and 
those with young zygospores show as bright a green as vigorously 
growing material. 

Spirogyra is not easy to keep in the laboratory. The small 
species keep better than the larger ones. Put only a small amount of 
the material in a jar and use rain water. If it is necessary to use tap 
water, let the water run for a minute before taking the water for the 
culture. Most metals are poisonous to Spirogyra, even the small 























186 


METHODS IN PLANT HISTOLOGY 


amount taken up by the water while standing in the water pipe being 
detrimental. 

The species found in running water will usually conjugate within 
a week when brought into the laboratory and placed in rain water or 
tap water. Species belonging to quiet waters, when brought into 
the laboratory and placed in a 0.2 per cent Knop’s solution, are 
likely to undergo rapid cell division and growth. After the alga 
has remained in such a culture for a few days or for a week, conjuga¬ 
tion may be induced by transferring to rain water or tap water, and 



Fig. 40.— Scenedesmus: photomicrograph from a preparation by Dr. Yamanouchi, mounted 
whole and stained as described in the text; Cramer contrast plate; 4-mm. objective; ocular X4; 
yellowish-green filter; camera bellows, 1 meter; arc light; exposure, 6 seconds. X675. 

keeping the culture in bright sunlight. Conjugation may begin 
within 3 or 4 days. Variations in temperature between 1° and 15° C. 
have little influence upon conjugation. 

The special chromo-acetic-osmic solution fixes well. Stain some 
material in iron-alum haematoxylin and some in Magdala red and 
anilin blue. Use the Venetian turpentine method, and on each slide 
mount material stained in both ways. With Magdala red and anilin 
blue the spiral chromatophore takes the blue and its pyrenoids the 
red. If the material contains figures, stain in iron-haematoxylin. 
This will stain the figures, but will hardly touch the chromatophore 




CHLOROPHYCEAE 


187 


or cell wall, thus allowing an unobstructed view of the figures. 
While figures occur occasionally in the daytime, collect your material 
at night, preferably near midnight. 

Spirogyra is easily imbedded and cut. 

Scenedesmus. —Scenedesmus (Fig. 40) is found everywhere as a 
constituent of the fresh-water plankton. It is more abundant in 
stagnant water. It often appears in considerable quantity in 
laboratory cultures, where it may be kept for years in a tightly closed 
glass jar without renewing the water, the lid being removed only 
when material is needed. 

The form is so small that in living material little more than the 
general form can be distinguished. Excellent mounts are easily and 
quickly made. Smear a very thin layer of albumen fixative upon the 
slide, and add a drop of water containing the Scenedesmus. The 
drop may be inverted for 1 or 2 minutes over the fumes of 1 per cent 
osmic acid. No washing is necessary, and good mounts may be 
made without any fixing whatever. Allow the drop to dry com¬ 
pletely. It is better to leave it for 24 hours before proceeding. The 
usual difficulty with this form, and with many others, is that the 
background stains and so makes the mounts untidy. The follow¬ 
ing method by Yamanouchi will produce beautiful preparations 
(Fig. 40): 

1. Dry on the slide, 24 hours. 

2. 10 per cent alcohol over night to remove chlorophyll. 

3. Safranin (alcoholic), 4 days. 

4. Water, 5 minutes. 

5. Aqueous gentian-violet, 2 days. 

6. Water, a few seconds. 

7. Orange G, aqueous, 3 minutes. 

8. 95 per cent alcohol, a few seconds. 

9. Absolute alcohol, 1 minute. 

10. Clove oil, until the stain is satisfactory. Different collections of 
Scenedesmus stain very differently, but the time in clove oil is likely 
to be long, even as long as 6 hours. 

11. Xylol, 5 minutes. 

12. Mount in balsam. 

Hydrodictyon.—-This is popularly known as the “water-net.” 
Hydrodictyon is found floating or suspended in ponds, lakes, or slow 
streams. The young nets are formed within the segments of the 


188 


METHODS IN PLANT HISTOLOGY 


older nets. Examine segments 4 or 5 mm. in length for the formation 
of young nets. The old nets may reach a length of 10 cm. Cultures 
are easily kept in the laboratory. If material which has been growing 
in a 0.5 to 1 per cent Knop’s solution be brought into tap water or 
pond water, zoospore formation may begin within 24 hours. Nets 
brought from the nutrient solution into a 1 to 4 per cent cane-sugar 

solution produce zoospores for 

mu Yi JJX 





m 






Jm 




bmv. 

c ? Tl 






a few days. 

Nets of all sizes should be 
selected for study. The seg¬ 
ments are coenocytic, and the 
nuclei of the older segments are 
hard to differentiate, except in 
stained preparations. Only one 
nucleus will be found in the 
young segments, but in the 
older segments the nuclei be¬ 
come very numerous. 

For fixing, use the special 
chromo-acetic-osmic formula. 
This should not produce plas- 
molysis in nets of any age. 
Iron-alum haematoxylin will 
differentiate the nuclei and 
pyrenoids, which may look 
alike with less precise stains. 
Use the Venetian turpentine 
method for mounting whole 
young nets and parts of older 
nets. Fine scissors should be used freely, because any attempt to 
arrange the material with needles will make it look as if the whole 
method of preparation were wrong. Parts of nets mounted whole are 
shown in Figure 41. 

For details of the formation of starch and for the finer details of 
the development of zoospores and gametes, Hydrodictyon should be 
imbedded and cut. 

Pleurococcus.—’This form, which is used everywhere as a labora¬ 
tory type of the unicellular green algae, is found on the bark of trees, 
where it is more abundant on the north side and near the ground. 


Fig. 41.— Hydrodictyon: A, part of young net 
with segments showing one pyrenoid and one or two 
nuclei; B, parts of three segments with nucleus, n, 
and pyrenoid, p, well differentiated; C, part of a still 
older segment with nuclei more numerous than the 
pyrenoids; D, part of a nearly mature segment with 
nuclei much more numerous than the pyrenoids. 
Fixed in the special chromo-acetic-osmic solution 
and stained in iron-alum haematoxylin. X600. 



CHLOROPHYCEAE 


189 


It is also found on stones and fences, and in moist situations generally. 
It is easily secured in nearly all localities and at all seasons. 

The life-history of Pleurococcus is variously described in textbooks, 
but it is very doubtful whether there is any mode of reproduction 
except by cell division. The zoospores and gametes which are some¬ 
times described probably belong to other forms which are occasionally 
associated with Pleurococcus, especially when growing in very moist 
situations. The life-history was ex¬ 
amined very critically by the great 
algologist, Wille, who not only con¬ 
cluded that cell division is the sole 
mode of reproduction, but showed how 
investigators, even those relying upon 
cultures, had made their mistakes. 

Wille’s paper was published in 1913 in 
Nyt Magazin for Naturvidenskaberne. 

A study of the living material is 
sufficient for any general course. The 
bright-green cells, scraped off and 
mounted in a drop of water, show the 
rather thick wall, the chromatophores, 
and usually the nucleus. A drop of 
iodine will bring out the nucleus, if it 
does not show already, and will also 
stain the pyrenoid, if the cell contains 
one. A mount in Venetian turpentine, 
stained in Magdala red and anilin blue, 
shows the nucleus very clearly. 

Vaucheria.—-This form can always 
be obtained in greenhouses, especially 
in the fernery, where it forms a green 
felt on the pots. The greenhouse 
form is likely to be Vaucheria sessilis. 

Another species, V. geminata, is very common in the spring, when it 
may be found in ponds and ditches (Fig. 42). Vaucheria is also found 
in running water, but in this situation is almost certain to be sterile. 
In the vicinity of Chicago, V. geminata appears late in March or 
early in April and within a few weeks begins to fruit abundantly. 
The fruiting continues for 4 to 8 weeks, and then the alga may 



Fig. 42.— Vaucheria: A, Vaucheria 
geminata, showing antheridium and five 
oogonia containing fertilized eggs; from a 
preparation fixed in formalin, acetic acid, 
and stained in iron-alum haematoxylin; 
B, V. sessilis ; from a preparation fixed 
in chromo-acetic acid and stained in eosin 
and gentian-violet. X150. 






190 


METHODS IN PLANT HISTOLOGY 


disappear until later in the season, when some of the oospores 
germinate. 

Vaucheria sessilis is found at all seasons in the greenhouses, but 
it is usually in the vegetative condition. Klebs found that the 
formation of oogonia and antheridia can be induced in V. repens 
(a variety of V. sessilis) within 4 or 5 days by putting the material 
into a 2 to 4 per cent cane-sugar solution in bright sunlight. The sex 
organs will not be formed in weak light or in darkness. 

The formation of zoospores may be induced in the following way: 
Cultivate in a 0.1 to 0.2 per cent Knop’s solution for a week, then 
bring the material into tap water, and keep the culture in the dark. 
Zoospores may appear within 2 days. Bright light or a temperature 
higher than 15° C. will check the production of zoospores. A 2 per 
cent cane-sugar solution kept in the dark is also likely to furnish 
zoosporic material. If no zoospores are formed when the solution 
is kept in the dark, the nutrition has been too weak: strengthen the 
nutrient solution and keep the culture in the light for a few days; 
then put the culture in the dark, and zoospores should appear. The 
formation of zoospores may continue for a couple of weeks. 

Aplanospore3 of V. geminata are formed in nature when the plant 
is growing upon damp ground. The aplanospores may also appear in 
a 4 per cent cane-sugar solution. 

In fresh 0.5 per cent Knop’s solution in bright light, cultures 
remain in the vegetative condition, and the result is the same in weak 
light if the nutrient solutions are seldom changed. Such cultures 
may be kept indefinitely by changing the nutrient solution whenever 
a whitish scum appears on the surface. 

Vaucheria is not easy to fix. Solutions which give fine results 
with Spirogyra and Zygnema may be ruinous to Vaucheria. We 
have secured the best results with a formalin-acetic solution (10 c.c. 
formalin, 5 c.c. glacial acetic acid, and 90 c.c. water). Chromic-acid 
solutions, even with 4 or 5 per cent acetic acid, cause some plasmolysis. 
If the chromic acid is weakened enough to prevent plasmolysis, the 
fixing is not thorough enough to prevent shrinking during subsequent 
processes. 

Iron-alum haematoxylin is the best stain. Magdala red and 
anilin blue give beautiful results, occasionally , but preparations are 
almost sure to fade. Eosin is good for topography, but will not show 
the nuclei. 


CHLOROPHYCEAE 


191 


Use the Venetian turpentine method. In mounting, use small 
scissors freely. You cannot untangle a mat of Vaucheria so as to 
give good views. 

For the development of the oogonium and antheridium, for 
fertilization and for the structure and development of the various 
spores, thin sections are necessary. Imbed in paraffin. For nuclear 
details, use iron-haematoxylin; for cytoplasm, 
use safranin, gentian-violet, orange. 

Cladophora.—'This genus is found in both 
salt and fresh water. The fresh-water forms 
are usually attached to sticks or stones in quiet 
or running water. The mats feel rough and 
crisp and, even under a pocket lens, show the 
characteristic branching by which the form is 
easily recognized. The absence of a mucous coat 
makes Cladophora a convenient host for numer¬ 
ous parasitic algae, among which diatoms be¬ 
longing to the genera Cocconeis and Gomphonema 
are particularly abundant. 

For laboratory cultures, select the forms 
found in quiet water, but for preparations, forms 
growing where the waves dash hard are better, 
since you can get a fine display of branches 
under a small cover. Forms growing in still 
water or in gently flowing water may look like 
unbranched filaments under an ordinary cover. 

The special chromo-acetic-osmic solution is excellent for Cladophora. 
Iron-alum haematoxylin, followed by the Venetian turpentine 
methods, gives the best results for nuclei and pyrenoids. Magdala 
red and anilin blue are better for the cell wall and chromatophores 
(Fig. 43). 

Ulothrix.—Where the problem of the origin and evolution of sex 
is studied, Ulothrix is an indispensable type. Ulothrix zonata is 
found in springs, brooks, and rivers, occurring in bright-green masses 
attached to stones in riffles, especially in sunny places. It is abun¬ 
dant on stones and piles along the beaches of lakes. Nuclear division 
takes place at night, most abundantly about midnight, and is followed 
by a rapid development of zoospores and gametes, which continue 
to be discharged throughout the forenoon. In the afternoon the 



Fig. 43.— Cladophora: 
fixed in chromo-acetic acid 
and stained in iron-alum 
haematoxylin. 












192 


METHODS IN PLANT HISTOLOGY 


material is largely vegetative. Another species is found in stagnant 
ponds, ditches, and even in watering-troughs and rain-barrels. It is 
difficult to keep in the laboratory the forms which are found in rapidly 
flowing water. However, if they are brought in still attached to 
stones and placed under a stream of tap water, they may live for a 
couple of weeks and may produce zoospores every morning. The 
production of zoospores may continue for a few days, if the material 
is merely put into a jar of water; in a 2 to 4 per cent cane-sugar 
'solution the production of zoospores continues a little longer. 

No form is better than Ulothrix for illustrating to a class the 
difference between zoospores and gametes. Even when gametes 
are not conjugating, their more rapid movement is noticeable; 
and when conjugating, the awkward, jerky movements of the pair 
contrasts sharply with the graceful movements of the zoospores. 

Fix in the special chromo-acetic-osmic solution and stain some 
material in iron-alum haematoxylin, which will differentiate the nuclei 
and pyrenoids; and stain some in Magdala red and anilin blue, 
which is better for the cell wall and chromatophore. Mount on 
each slide material from both lots, and the preparation will then 
afford a rather complete study. 

Oedogonium.—Most species are found in quiet waters, especially 
in ponds and ditches. The best fruiting material is often attached to 
submerged twigs, rushes, and various plants, where, to the naked eye, 
it forms only a fuzzy covering. Some species form floating masses, 
bearing some resemblance to Spirogyra, but they are not so slippery. 

The special chromo-acetic-osmic solution is good for the filaments; 
the iodine solution used in testing for starch, or a weak aqueous 
solution of osmic acid (4 or 5 drops to 50 c.c. of water) is good for 
zoospores and androspores. 

For cell division and the peculiar method of forming the new cell 
wall, stain in Magdala red and anilin blue. Iron-alum haematoxylin 
is better for most of the other phases; but the fertilized eggs stain 
very deeply. Consequently, stain some material lightly, for the 
fertilized eggs; and some more deeply for young eggs, chromatophores, 
and other phases. Mount in Venetian turpentine. 

For details of blepharoplasts and the development of the various 
motile forms, material should be imbedded and sectioned. 

Nanandrous species have antheridia only in the dwarf males; 
and species with antheridia in the ordinary filaments have no andro- 


CHLOROPHYCEAE 


193 


spores or dwarf males in the life-history. A species with dwarf 
males is shown in Figure 44. 

In studying Oedogonium diplandrum, Klebs found that a change 
from a lower to a higher temperature would 
induce the production of zoospores. A culture 
which had been kept in a cold room with a 
temperature varying from 6° to 0° C., when 
brought into a warmer room with a tempera¬ 
ture varying from 12° to 16° C., produced an 
abundance of zoospores within 2 days. Light 
does not seem to have any influence upon the 
formation of zoospores in this species, but 
light is necessary for the formation of anther- 
idia and oogonia. 

We have secured an abundance of oogonia 
and antheridia by keeping the material for 4 
or 5 days in a very weak Knopf’s solution 
and then tranferring to distilled water. The 
oogonia appeared in 3 or 4 days. The method 
seems to succeed with some species, especially 
those which occur floating or suspended in the 
water, but we have not succeeded with species 
which form a fuzzy covering on grasses and 
twigs under water. Sterile material some¬ 
times fruits when brought into the laboratory 
and placed in open jars with plenty of water 
and not too much light. 

Coleochaete. — Coleochaete is epiphytic upon 
the stems and leaves of submerged plants. 

C. scutata, which is the most common species, 
has a flat, orbicular thallus generally less than 
1 mm. in diameter. C. pulvinata has a hemi¬ 
spherical thallus and might be mistaken for 
Rivularia, unless examined with a lens. 

For most purposes it is better to mount 
the whole plant, in situ on the leaf upon which 
it is growing. If it is growing on stems or 
petioles, strip off the epidermis and mount small pieces. In this way 
you get the very young stages, from the fertilized egg up to the adult 



Fig. 44. — Oedogonium: 
an undescribed species col¬ 
lected by Dr. Elda Walker. 
In A, on the left, is a dwarf 
male still showing the cap cell; 
on the right, the cap cell has 
come off and the antheridium 
at the top has discharged one 
of the sperms, but the other, s, 
still remains; in the anther¬ 
idium below are two young 
sperms. B, upper part of a 
cell, showing the cellulose ring 
which will form the side wall 
of the new cell. The nucleus 
is in the spirem stage of 
division. X 280. 










194 


METHODS IN PLANT HISTOLOGY 


plant. The young stages are likely to be overlooked and even when 
noticed are likely to be lost if you attempt to remove Coleochaete 
from its host. If you think you must remove it from the host, 
complete the staining before the removal. 

Fix in the special chromo-acetic-osmic solution and stain sharply 
in Delafield’s haematoxylin. If you overstain and reduce with 
hydrochloric acid (3 or 4 drops in 100 c.c. of water), the Coleochaete 
will stand out against the host, which is not likely to stain so deeply 
as the alga. A slight tinge with orange in clove oil may increase the 
contrast between the alga and its host. 

After staining, the material may be run up through a series of 
alcohols, 5, 10, 20, 35, 50, 70, 85, 95, and 100 per cent, about 15 
minutes in each grade. If orange in clove oil is used, apply it here 
and transfer to xylol. Mount in balsam. If no orange is used, 
transfer to 25, 50, 75, and 100 per cent xylol, and mount in balsam. 
After staining, the Venetian turpentine method may be used instead 
of the rather long series of reagents. 

Sections are easily cut and, especially in forms with a flat thallus, 
show features which might escape if one depended entirely upon 
plants mounted whole. Cut out small pieces of leaf or stem 
abundantly covered with Coleochaete , imbed in paraffin, and cut 
host and guest together. 

Chara.— Chara is found in ponds, lagoons, and ditches. Once 
seen, it is always readily recognized. In the ponds and lagoons 
along the southern shores of Lake Michigan it fruits so abundantly 
that the whole pond shows an orange color due to the immense 
numbers of antheridia. In the lagoons of the Chicago parks Chara 
is so abundant that it must be dredged out every summer. 

Chara is easily kept alive throughout the year in the laboratory. 
A 2-gallon glass jar with an inch of pond dirt, sand, and gravel at 
the bottom, and nearly filled with tap water, is all that is needed for 
a successful culture. If the jar is to be covered, it should not be 
more than two-thirds full of water. Not more than a dozen plants 
should be put into such a jar. 

A rather strong solution should be used for fixing. The following 


will give good results: 

Chromic acid . 1 g. 

Glacial acetic acid. lc.c. 

Water. .. 100 c.c. 





CHLOROPHYCEAE 


195 


In about 24 hours this not only fixes, but it dissolves the lime with 
which most species are coated. 

For paraffin sections select the tip of the plant, a piece about 
a centimeter in length. Sections of this may show, no,t only the large 





Fig. 45.— Chara: fixed in chromo-acetic acid and stained in safranin, gentian-violet, orange. 
A, longitudinal section of apex showing five nodes; c, corticating filaments; n, part of the frag¬ 
mented nucleus. B, note the different condition of the nodes and first branches. C, young antherid- 
ium, o, and young oogonium, o. X230. 


apical cell, but also various stages in the development of antheridia 
and oogonia (Fig. 45). For the development of the plant body from 
the apical cell and also for early stages in the development of oogonia 
and antheridia, the safranin, gentian-violet, orange combination is 


















196 


METHODS IN PLANT HISTOLOGY 


excellent; for later stages, especially in the development of the 
antheridia, iron-haematoxylin is much better. 

The antheridium of Chara stains so rapidly that the beginner uni¬ 
formly makes poor preparations. In order to get good preparations 


CHLOROPHYCEAE 


SIPHONEAE 

Coenoctjtic 
Thij/losipbonaceae 

I 

i 

i/auc/ji 


'auchenaceae 


fophomceae 


/ CONFERVOIDME 

Filamentous, uninucleate 

Co!co chaetaceac 


Fterbesiacecye 
Cauleifaceac Codiaccae 

Bruopd/daceae 

' | JSasyc/adaceae 
j /'.lA/oniaceae 
Bot ry cliia cca e 

'//t/drodictifon 
v „ ocenedetmus 

\ M/m . 

Cudonna 
/' qL rnndorina 

o u c Q ^ Ullamaaomonas 


i 

Chaetoporaceae 


v „ ..-yjoniarm 

Ulotricfi'aceae 


U/i 


vat'eae 


P/eurococcus 


^protococcoideae; , , 

[lot ft lamentous, mostly uninucleate 

Fig. 46.—A diagram illustrating an opinion in regard to relationships in the Chlorophyceae 


of the antheridium, it is necessary to disregard other structures, 
which will be stained lightly or not at all when the stain is just right 
in the antheridial filaments. 



CHLOROPHYCEAE 


197 


If it is desired to mount whole branches showing the antheridium 
and oogonium in position, use the Venetian turpentine method, 
staining in Magdala red alone, or in Magdala red and anilin blue. 
Good mounts showing shield, manubrium, capitula, and filaments 
may be obtained by crushing an antheridium under a cover-glass. 
For this it is better to stain in Magdala red alone, since any over¬ 
staining is easily corrected by exposing the preparation to direct 
sunlight. 

As slides accumulate, the thoughtful student will feel the need 
of some kind of classification. Of course, one might arrange alpha¬ 
betically and there would be no need for thought; but we assume 
that the student who has made enough slides to need a classification 
will want one which expresses some idea of relationship, even if 
the idea may be more or less faulty. The classification indicated 
in Figure 46 is essentially that of Oltmann, and does not differ 
much from that given in Engler and Prantl’s Die natiirlichen Pflan- 
zenfamilien. If the student would compile similar diagrams in all 
the groups, his slides would mean, not only proficiency in technic, 
but an increasing knowledge of the structure, development, and 
relationship of plants. 


CHAPTER XVI 

PHAEOPHYCEAE. BROWN ALGAE 

The Phaeophyceae, or brown algae, are almost exclusively 
marine. They include a great variety of forms, ranging from delicate 
filaments to coarse, leathery plants 100 feet in length. There 
are no unicellular members. 

For fixing marine algae, fixing agents should be made up with 
sea-water, never with fresh water, and the washing should be done 
with sea-water; but fresh water should be used in making the series of 
alcohols. When the Venetian turpentine method is to be used, wash 
in fresh water before placing in the 10 per cent glycerin.^ 

For habit work, material may be put into formalin—about 6 c.c. 
commercial formalin to 100 c.c. of sea-water—and kept there indefi¬ 
nitely. If it is desired to transport large quantities of coarse forms, 
the material may remain in this solution for a week and may then be 
removed from the liquid and packed in closed pails or tubs or any 
water-tight containers. After reaching its destination, the material 
should be put into formalin again. 

For material to be mounted by the Venetian turpentine method, 
6 to 10 per cent formalin (always in sea-water) is a good fixing agent. 
Wash in sea-water for 1 hour, then in equal parts sea-water and fresh 
water for J hour, then in fresh water | hour. The material is now 
ready for staining in aqueous stains, or for the 10 per cent glycerin, 
if alcoholic stains are to be used. 

The following formula by Flemming will also give good results, 
both for the Venetian turpentine method and for the paraffin method: 


Chromic acid . 1 g. 

Glacial acetic acid. 0.4 c.c. 

Sea-water. 400 c.c. 


Fix 24 to 48 hours and wash 24 hours in running sea-water. A 
convenient washing-box can be made from an ordinary washtub. 
Bore a dozen f-inch holes in the bottom; insert rubber tubes 
6 inches long, and in the end of each tube place the glass part of a 
pipette. The tub may be elevated by nailing three narrow boards 

198 





PHAEOPHYCEAE 


199 


to the sides so as to form a tripod. Place the bottles or cans of material 
under the pipettes and let sea-water flow into the tub. 

If such chromic-acid material is to be used at once for Venetian 
turpentine mounts, follow the washing in sea-water by \ hour’s 
washing in equal parts sea-water and fresh water (not necessarily 
running water) and then \ hour’s washing in fresh water. The 
material is now ready for an aqueous stain or for 10 per cent glycerin. 
If desirable to keep it for future staining, put it into 5 or 6 per cent 
formalin in fresh water. 

Material for sections may be treated in the same way, but it is 
often better to add 2 to 10 c.c. of 1 per cent osmic acid to 100 c.c. 
of the chromic-acid solution. The 1 per cent osmic acid should be 
made up in distilled water. 

Small filamentous forms, if they are to be mounted whole, are 
usually more satisfactory to handle if fixed in formalin. A strong 
solution, about 10 c.c. of commercial formalin to 90 c.c. of sea-water, 
is very good. Material is well fixed in a few days, but may be left 
here until it is needed. To make preparations, wash in fresh water 
and follow the Venetian turpentine method. 

For habit demonstrations many of the smaller forms can be 
floated out and dried on paper. Ectocarpus, Desmotrichum , Dictyota, 
Cutleria, and even small specimens of Laminaria are quite useful 
when prepared in this way. Take a light pine board, a little larger 
than the standard herbarium sheet, float it in a tub of water, place 
on the board the paper upon which the material is. to be mounted, 
arrange the material with a toothpick or the blunt end of a needle, 
dipping all or a part of the board under water whenever necessary. 
Cover with a piece of cheese-cloth, add a blotter or two, as in case 
of flowering plants, and dry under gentle pressure, changing the 
blotters frequently. The algae have enough mucilage to make them 
adhere to the paper. Coarse forms, like Fucus , may need to be held 
down by strips of gummed paper. 

Sphacelaria.—-The apical cell of Sphacelaria or the nearly related 
Stypocaulon affords an excellent study of the structure of cytoplasm. 
Flemming’s weaker solution, with the osmic acid even a little weaker 
than recommended in the formula, is good for the apical cell and the 
mitotic figures, which are quite conspicuous. For these features 
it is a good plan to break off the tips so as to have only pieces 6 to 
12 mm. long, which will lie flat in the paraffin. The tips should be 


200 


METHODS IN PLANT HISTOLOGY 


broken off after the material has been brought into xylol. If 
whole tufts are imbedded, the branches diverge enough to make 
perfectly longitudinal sections of the apical cells rather rare. Iron 
haematoxylin with a faint staining in orange is a satisfactory com¬ 
bination. 

Ectocarpus.—For general morphological study, branches should 
be mounted whole in Venetian turpentine. A 6 to 10 per cent 

formalin solution (in 
sea-water, of course), 
or the chromo-acetic 
acid will give good fix¬ 
ation. Stain some in 
iron-haematoxylin and 
some in Magdala red 
and anilin blue. 
Mount on each slide 
some from each lot. 
Sporangia usually ap¬ 
pear earlier than the 
gametangia. So col¬ 
lections should be 
made at different sea¬ 
sons. August is rather 
late for sporangia, but 
gametangia are abun¬ 
dant at this time. You 
should have both spor¬ 
angia and gametangia 
on each slide (Fig. 47). 

Desmotrichum.—Forms as large as Desmotrichum can be handled 
like Ectocarpus , but care must be taken not to overstain. 

Laminaria.—-In such large forms, small portions showing the 
structure and development of the thallus and also the reproduction 
should be cut out with a razor and then placed in the fixing agent. 
The sporangia of Laminaria stain very deeply and quickly. Iron- 
haematoxylin is good, but be careful not to overstain. After this 
stain is just right, about 3 to 5 minutes in alcoholic safranin will 
stain the mucilaginous structures and add to the value of the 
preparation. 




Fig 47.— Ectocarpus: A, sporangia (“unilocular spor¬ 
angia”) of various ages; gametangia (“pleurilocular gametan¬ 
gia”), three of them nearly mature. Fixed in 10 per cent 
formalin and stained in iron-haematoxylin. X280. 









PHAEOPHYCEAE 


201 


The zoospores from the sporangia germinate immediately, 
forming dioecious, filamentous gametophytes bearing antheridia and 
oogonia. The fertilized egg at once begins to develop into the 
Laminaria plant. So little is known about this stage that it would 
be worth while for those who are at the seashore to germinate the 
zoospores of any members of the Laminariaceae. 

For habit study, small specimens up to 45 cm. in length can be 
mounted upon paper. They stick well and seldom need to be secured 
by gummed paper. Larger specimens may be allowed to dry and 
may then be stored away in a box. When wanted for use, wet them 
under the tap, or, better, 
in salt water; after using, 
let them dry and return 
them to the box. Speci¬ 
mens will stand 4 to 5 
such soakings in fresh 
water; if a pint of salt is 
added to 3 or 4 gallons of 
water, the material may 
be soaked a dozen times 
before it passes its useful¬ 
ness. If material has 
been fixed in formalin, it 
may be washed in sea¬ 
water—not very thor¬ 
oughly, but enough to 
remove the pungent odor 
—and then soaked in 

equal parts of glycerin and water. Use only enough of the glycerin 
to make the specimens flexible, not enough to make them wet to 
handle. In this way, material of Laminaria , Macrocystis, Nereocystis, 
Postelsia, and other large forms can be kept in condition for demon¬ 
stration and will last for years without any attention. When not 
in use, they should be kept stored in a box. 

Cutleria. —This alga deserves a place in any course in morphology, 
if the course is thorough enough to permit the study of three members 
of the Phaeophyceae. These three should be Ectocarpus (or 
Pylaiella), Cutleria , and Fucus. Cutleria is not found on the Ameri¬ 
can coasts, but is abundant at Naples. The habits of gametophyte 




Fig. 48 .—Cutleria multifida: A, oogonia; B, an¬ 
theridia. Fixed in Flemming’s weaker solution, cut 3/i. 
and stained in iron-alum haematoxylin. X470. 




202 


METHODS IN PLANT HISTOLOGY 


(known as Cuileria) and the sporophyte (known as Aglaozonia) are 
so different that they furnish a good illustration of alternation of 
generations. Beginners understand such an illustration more readily 
than they do an illustration like Didyota, with its two generations 
looking so nearly alike. Cutleria also furnishes a good stage in the 
evolution of sex, about midway between isogamy and the extreme 
heterogamy of Fucus. 



Fig. 49. —Fucus vesiculosus: A, antheridial branch with antheridia in various stages of 
development; B, the third mitosis in the antheridium, one figure showing a transverse view in 
which the chromosomes can be counted; C, the fourth mitosis, with two of the eight figures cut 
transversely but hard to count chromosomes in the figure, which is reduced one-half; D, the 
third mitosis in the oogonium, showing centrosomes and radiations. Fixed in Flemming’s weaker 
solution, cut 3/x, stained in iron-alum haematoxylin. X480. 

For habit study, both generations should be mounted upon paper. 
The gametophyte ( Cutleria ) sticks well, but the sporophyte ( Agla¬ 
ozonia ) will need some glue or gummed paper. 

For paraffin sections, fix in chromo-acetic acid. Cut 10 y thick. 
For mitotic figures, some osmic acid should be added to the chromo- 
acetic acid and the sections should be much thinner, about 3 to 5 /x- 
Use iron-haematoxylin and then stain for 3 to 5 minutes in alcoholic 
safranin (Fig. 48). 

Fucus. —Material for habit study may be dried, or preserved in 
formalin, or mounted on paper. In the latter case, glue or gummed 
paper will be necessary. Most satisfactory of all is to send to Woods 




PHAEOPHYCEAE 


203 


Hole, Massachusetts (George M. Gray), for living material. Fertili¬ 
zation occurs at all seasons, but autumn is the most favorable. In 
summer the material dies before it reaches Chicago, but during the 
rest of the year a pailful will reach Chicago, and even as far west as 
the Mississippi River, in good condition for showing the rotation 
of the egg by the sperms. The eggs and sperms form slimy masses, 
the antheridia being orange red, and that containing the eggs a dirty 
green. Mix a drop of the red with a drop of the green. The move¬ 
ments of the egg can be observed, and material for a study of fertili¬ 
zation and later stages is easily secured. In fixing fertilization 
and succeeding stages, it is worth while to use some of the regular 



Fig. 50. —Dictyota dichotoma: longitudinal section through apical region showing two apical 
cells, the one on the right dividing again to produce another dichotomy. Stained in iron-alum 
haematoxylin. Photomicrograph by Miss Ethel Thomas. X167. 


Flemming's weaker solution, as well as the solution without the 
osmic acid. 

For the growing points and conceptacles, small pieces should be 
cut off with a razor. If the fruiting tips be cut through lengthwise 
before they are cut off, the fixing will be more satisfactory. For 
sections of the conceptacles it is better not to cut across the whole 
tip, but to cut off pieces of the rind containing half-a-dozen concep¬ 
tacles. Such pieces are more easily imbedded and cut. There is no 
difficulty in cutting such pieces in paraffin. Iron-haematoxylin is a 
good stain. Safranin and gentian-violet are also satisfactory, but 
care must be taken not to overstain since Fucus usually stains deeply 
and rapidly. 

For the cytologist, Fucus might be used as a test object for testing 
proficiency in technic, just as Pleurosigiua angulatuTn is used in test¬ 
ing an objective. The nuclear divisions in the antheridium are 




204 


METHODS IN PLANT HISTOLOGY 


simultaneous, and at the sixth division, which is the last, there are 
32 mitotic figures, each with 32 chromosomes which split so that 32 
go to each pole. When you can make a preparation in which these 
chromosomes can be counted, your technic is adequate for research 
work in cytology. In a good preparation, the mitotic figures in the 
oogonium show small but brilliant centrosomes, with a great display 
of radiations. (Fig. 49). 

Dictyota .—Didyota deserves a place in the series illustrating the 
evolution of sex, since its large egg has lost all motility, but the 
difference in size of egg and sperm is not so extreme as in Fucus. 
It also furnishes an excellent example of the development of a thallus 
from an apical cell (Fig. 50). 

Mount habit material on paper. For sections, fix in chromo- 
acetic acid. For figures, cut 3 to 5 y, but for general views of apical 
cell and reproductive phases, cut 10 y. Stain in iron-haematoxylin 
and counter-stain for 2 or 3 minutes in safranin. 


CHAPTER XVII 

RHODOPHYCEAE. RED ALGAE 

The red algae belong almost exclusively to salt water, but a few 
genera are found only in fresh water, usually in running water, and 
a few forms occur both in salt and in fresh water. Nearly all are 
small forms, and for habit work can be floated out and mounted 
on paper. Very few will need glue or gummed paper. 

For more critical habit work and for Venetian turpentine mounts, 
fix in 6 to 10 per cent formalin in sea-water. Material keeps indefi¬ 
nitely in 10 per cent formalin. 

For sections, use the chromo-acetic acid with or without the addi¬ 
tion of a little osmic acid, as recommended for the brown algae. 
The same method of fixing and washing should be used as for the 
brown algae, except that in the case of the few fresh-water forms, fresh 
water should be used in making the fixing agent and in washing it out. 
For Polysiphonia, and doubtless for many other forms, the period 
in the fixing agent should be very much shortened. Picric acid, 
corrosive sublimate, and absolute alcohol have been tried, but the 
results have not been encouraging. 

Batrachospermum.—-This is a green, fresh-water member of the 
red algae. It is not very uncommon in small streams. Fix in 
chromo-acetic acid (in fresh water) and use the Venetian turpentine 
method. Good preparations showing the nuclei may be obtained by 
staining in Mayer’s haem-alum, or Haidenhain’s iron-haematoxylin. 
After the material is ready for mounting, tease out a small portion, 
and still further dissociate the filaments by tapping smartly on the 
cover. 

Nemalion.—Methods for preparing Nemalion have been described 
by Wolfe. Chromo-acetic acid proved to be most satisfactory for 
fixing. For studying fertilization, mounts were made as follows: 

Young tips were crushed in water under a cover-glass and on a slide 
that had previously been treated with fixative; the cover was then removed, 
and the water on the slide ‘allowed to evaporate. The gelatinous nature of 
the wall prevents the contents of the cell from being affected by this treatment, 

205 


206 


METHODS IN PLANT HISTOLOGY 


even when the albumen has hardened sufficiently to hold the filaments firmly 
in place. 1 

Stain in safranin and gentian-violet, and mount in balsam. 

Iron-haematoxylin is recommended for paraffin sections. The sec¬ 
tions must be very thin, 5 /jl or less. “ Material killed in 2 per cent 
formalin in sea-water and gradually transferred to pure glycerin kept 
its color perfectly.” 

For material to be mounted whole, we should recommend fixing 
in 10 per cent formalin and staining in iron-alum haematoxylin. Place 



Fig. 51 .—Nemalion multifidum: A, branch showing carpogonium, c; trichogyne, t; central 
strand, m. B, somewhat older stage showing two-cell stage immediately following fertilization, e; 
carpogonial branch, b. C, branch with antheridia, a. Fixed in 10 per cent formalin and stained 
in iron-alum haematoxylin. X400. 

the material in 10 per cent glycerin until all the water is out. Mount 
in glycerin j elly. To make a mount, take a small piece of the material, 
not more than 3 or 4 mm. long, touch it to filter paper to remove as 
much of the glycerin as possible, put it into the melted glycerin 
jelly, add a round cover, and crush by tapping on the cover. The 
antheridia and procarps are in the slender filaments, and the cysto- 
carps are in the larger filaments. If several slides are to be made, 
it is a good plan to select slender, medium, and thicker filaments, 

1 James J. Wolfe, “Cytological Studies in Nemalion,” Annals of Botany, 18-607- 
630, 1904. 





RHODOPHYCEAE 


207 


remove the surface glycerin with filter paper, and then crush the fila¬ 
ments between two slides. There is scarcely any danger of crushing 
too much. A little of the crushed material, including the various 
stages, can be put into the melted glycerin jelly. Add a round cover, 
tap gently until the jelly comes just exactly to the edge of the cover. 
As soon as the jelly is cool, the mount may be sealed with balsam 
but we prefer to leave the mounts for a day or two before sealing. 
Such mounts would probably keep for a year or two without sealing 


(Fig. 51). 



Fig. 52 .—Polysiphonia fibrillosa: A, nearly mature cystocarp, showing the large cell formed 
by the fusion of several auxiliary cells with the pericentral cell—the carpospores are large and elon¬ 
gated; B, an antheridium—the term “ antheridium ” is more correctly applied to the structure shown 
in C, a, which cuts off one or more sperms, s; D, young tetraspores. Fixed in Flemming’s weaker 
solution, cut 3 n, and stained in iron-alum haematoxylin. A and B X240; C and D X780. 


We have mounted Nemalion in Venetian turpentine; but by 
this method the material becomes hard and behaves like cartilage, 
so that it cannot be crushed under a cover. However, it can be 
crushed on a piece of glass with a scalpel. 

Nemalion, stained in eosin, makes beautiful mounts, but they 
always fade. 

Polysiphonia.—This is a very difficult form to handle, but Dr. 
Yamanouchi has developed an adequate method, and, by following 
it, anyone should be able to get good preparations. 

For mounting in glycerin, glycerin jelly, or in Venetian turpentine, 
fix in 10 per cent formalin and stain in iron-haematoxylin. 



208 


METHODS IN PLANT HISTOLOGY 


For sections, fix in Flemming’s weaker solution, but omit the 
osmic acid for spermatogenesis and germination of carpospores. 
The time should be very short, 5 to 40 minutes being sufficient. If 
material is left too long, it goes to pieces. Wash in a gentle stream of 
sea-water for 24 hours. Stain in iron-haematoxylin and then for 
2 to 3 minutes in safranin (Fig. 52). This short stain in safranin 
gives a faint rosy tinge to mucilaginous structures, but does not obscure 
the fine nuclear detail. In the nucleus of the sperm, the chromosomes 
remain distinct, so that the number, 20, can be counted from the 
time the sperm is formed (Fig. 52C) up to fertilization. 

With very delicate forms, like Callithamnion and Griffithsia , the 
washing may be in part or even wholly omitted, and the chromic acid 
extracted by the lower alcohols, the material being kept in the dark. 

Corallina.— Corallina and other forms whose surface is incrusted 
with lime need special treatment. The following solution is good: 


Chromic acid. 1 g. 

Glacial acetic acid. 1 c.c. 

Sea-water. 100 c.c. 


Fix 24 hours, changing the fixing agent 2 or 3 times. Wash 24 
hours in sea-water. 

If carefully applied, the following is a good method: Put the 
material into 5 per cent glacial acetic acid (in sea-water) and watch 
it. As soon as the vigorous effervescence begins to subside, rinse 
in sea-water and transfer to Flemming’s weaker solution, and fix 
24 hours. Iron-haematoxylin is best for figures, but for general 
structure the safranin, gentian-violet, orange combination gives 
beautiful results. 





CHAPTER XVIII 


FUNGI 

In recent years such progress has been made in culture methods 
for fungi, especially for those which cause disease, that only the trained 
pathologist could be expected to know just what is the best medium 
for each particular fungus. We shall not attempt to deal with the 
subject of culture media, but shall simply indicate how a student, 
not trained in pathology, may secure material for preparations. 
Professor Kleb’s methods make it possible to secure material of 
many forms in various phases of their life-histories. 

In general, filamentous fungi are treated like the filamentous 
algae, while the fleshy forms are cut in paraffin. 

PHYCOMYCETES 

Rhizopus (Mucor).—This familiar mold appears with great 
regularity on bread. The following is a sure and rapid method for 
obtaining Mucor: Place a glass tumbler in a plate of water, put on 
the tumbler a slice of bread which has been exposed to the air for 
a day, and cover with a glass j ar. The bread must not become too wet. 

To obtain a series of stages in the development of the sporangium 
it is better to use living material. For class work, time the cultures 
so as to have a plenty of sporangia which have not yet begun to 
turn brown. 

For permanent preparations, fix for at least 24 hours in 10 per cent 
formalin; wash J hour in water, and then follow the Venetian turpen¬ 
tine method. Eosin gives good results. Stain over night or 24 
hours, treat with 2 per cent acetic acid, changing several times, and 
then put into glycerin, merely pouring off the 2 per cent acetic acid 
and not rinsing the acid out in water. When washing out the glycerin, 
do it with alcohol which has about 0.5 c.c. of acetic to the 100 c.c. 
of alcohol, and leave 1 or 2 c.c. of this slightly acid absolute alcohol 
on the material when you add the 10 per cent Venetian turpentine. 

For permanent preparations, fix for at least 24 hours in the special 
chromo-acetic-osmic solution; wash 24 hours in water and follow 

209 


210 


METHODS IN PLANT HISTOLOGY 


the Venetian turpentine method. Eosin gives good results for general 
topography. Stain over night or 24 hours, treat with 2 per cent acetic 
acid in water for 5 or 10 minutes, changing several times, and theri 
put the material into 10 per cent glycerin, merely pouring off the acid 
but not washing it out with water. Wash out the glycerin with 
alcohol containing about 0.5 c.c. acetic acid to 100 c.c. of the 95 and 
100 per cent alcohol. Leave about 1 c.c. of the slightly acid alcohol 
on the material as you add the 10 per cent Venetian turpentine. 

To bring out the nuclei, use iron-alum haematoxylin. Rhizopus 
stains very rapidly, so that an hour in iron-alum and an hour in the 
haematoxylin may be long enough. The sporangia stain more readily 
than the mycelium; consequently to show the ‘coenocytic character 
of the mycelium, the action of the second iron-alum must be stopped 
earlier than when staining for the sporangium. Extract the stain 
until the nuclei of the mycelium show clearly, and then remove part 
of the material to wash in water. For the rest of the material, 
continue to extract the stain until the sporangia are satisfactory. 
Mount some of each lot on each slide. 

The finer details of the sporangium can be seen only in thin sec¬ 
tions. Rhizopus is the most easily obtained material for showing the 
progressive cleavage of protoplasm by vacuoles. 

The zygosporic stage in the life-history is rarely met in nature 
or in cultures, but when once secured it may be propagated indefi¬ 
nitely. We have a culture which has been furnishing illustrative 
material for nearly twenty years. Once in a while, when a particu¬ 
larly good culture appears, lay aside some of it to start the next 
culture. The best series of stages generally appears between the 
fourth and seventh days. Dr. Blakeslee shows why zygospores are 
so infrequent. The conjugating filaments belong to different strains 
of mycelia which he calls “plus and minus strains,” and which, for 
convenience, may be called “female and male strains.” The more 
vigorous mycelium is +, and the less vigorous —. When the two 
strains come together, zygospores are formed along the line of meeting. 
If -f- and — strains are started at opposite sides of a dish, they will 
meet near the middle and form a dark line of zygospores. Through 
Dr. Blakeslee’s generous distribution of material, the + and — strains 
are now available in practically all of the great universities of the world. 

While Rhizopus may be grown on bread, it is better to use culture 
media in Petri dishes. While it grows well on agar media, it is hard 


FUNGI 


211 



to pick it off; and on liquid media, the growth is abnormal. Mrs. 
Alice A. Bailey has devised a method which is ideal for securing ma¬ 
terial. She puts about the usual amount of a potato dextrose 
agar in a Petri dish and pours over it a potato decoction about 2 mm. 
deep. The + and — strains are then added. The potato decoction 
is made as follows: Use 300 g. of Irish potatoes and 1000 c.c. distilled 
water. Peel and slice the potatoes and boil for 1 hour in the distilled 
water. Strain off the liquid through cheese-cloth and make up to 
the original quantity by 
adding distilled water. 

Flask and sterilize. 

The potato dextrose 
agar is made as follows: 
to 1,000 c.c. of potato 
decoction, add 20 g. of 
dextrose and 30 g. of 
agar. Boil | hour in 
a double boiler. Strain, 
flask, and sterilize. 

The potato dextrose 
agar is an excellent 
medium and the thin 
layer of potato decoction 
keeps the material from 
sticking to the agar so 
that it can be lifted off 
intact. Binse it under 
the tap and fix in 
the chromo-acetic-osmic 
solution. 

Zygospores may begin 
to form within 3 days, 
and mature zygospore xso. 
may appear within 4 or 

5 days. Watch the cultures and fix so as to secure a series of stages. 
(Fig. 53). 

Paraffin sections should not be thicker than 5 n, and 3 n is better 
for nuclear detail. Iron-alum haematoxylin is best for nuclei, but 
safranin, gentian-violet, orange will give beautiful preparations. 


Fia. 53 .—Rhizopus nigricans: various stages in the 
development of zygospores from a culture on bread; prepara¬ 
tion stained in eosin and mounted in Venetian turpentine. 






212 


METHODS IN PLANT HISTOLOGY 


In the related genus, Sporodinia, which is rather common in 
summer on fleshy fungi, especially upon Boletus and its allies, the 
zygosporic condition is not infrequent, because Sporodinia does not 
have + and — strains. Rhizopus behaves like a dioecious plant, 
while Sporodinia behaves like a monoecious one. The very damp 
atmosphere and the nutrition necessary for the formation, of zygo¬ 
spores may be provided in the laboratory in the following way: Put 
a little water in a glass battery jar and place filter paper around the 
inside of the jar so that it will take up water and thus keep the sides 
of the jar moist. Place a small beaker or dish, without any water in 
it, in the bottom of the jar, and in the beaker place a small piece of 
bread dampened with the juice of prunes. Infect the bread with 
spores, or use a piece of bread upon which mycelium is already grow¬ 
ing. Sections of the root of Daucus carota may be used instead of the 
bread. Put a piece of wet filter paper on a pane of glass and cover 
the jar. Begin to examine after 24 hours. The zygospores may 
appear in 4 or 5 days. A very full account of the methods by which 
the various phases of the life-history of Sporodinia may be produced 
at will is given by Klebs in the Jahrbiicher fiir wissenschaftliche 
Botanik, 32:1—69, 1898. 

Zygorhynchus is another interesting relative of Rhizopus, readily 
distinguished by having suspensors of very unequal size. Dr. 
Florence A. McCormick sent us magnificent zygosporic material, 
raised on beef broth and fixed in 10 per cent formalin in water. 

Saprolegnia.—This is an aquatic mold, very common upon 
insects and algae. Cultures are easily and quickly made. Bring 
in a quart of water from any stagnant pond or ditch, and into the 
water throw a few flies. After 12 to 24 hours throw the water away, 
rinse the flies in clean water, and put then into tap water. The 
water must be changed every day to keep bacteria from ruining the 
culture. The larvae of ants or small pieces of boiled white of egg are 
sometimes better than flies. Sporangia may appear within 24 hours 
but may be a day later. Sporangia may be produced in the greatest 
abundance by cultivating the mycelium for several days and then 
transferring it to pure water or to distilled water. As long as the 
nutrient solution is sufficiently strong and fresh, only sterile mycelium 
will be produced. 

To secure oosporic material, mycelium which has been highly 
nourished for several days in a nutrient solution is brought into a 


FUNGI 


213 


0.1 per cent solution of leucin, or into a 0.05 to 0.1 per cent solution of 
haemoglobin. Begin to examine after 24 hours. 

Oogonia have been produced in great numbers by the following 
method: cut ordinary corn (Zea Mais) into small pieces and boil for 
20 minutes. When cool, put pieces into a Petri dish and add enough 
pond water to nearly cover the pieces of corn. Oogonia may appear 
within 3 or 4 days. 

For fixing, the following formula is excellent for material which 
is to be mounted whole: 

Formalin. 10 c.c. 

Glacial acetic acid. 5 c.c. 

Water. 85 c.c. 

Fix at least 24 hours, but material may be left for months in this 
fixing agent. 

Stain some in Magdala red and 
anilin blue, and some in iron-alum 
haematoxylin. Mount some of each 
lot on each slide (Fig. 54). 

For sections, it is better to fix in 
the special chromo-acetic-osmic-acid 
solution. 

Satisfactory material for general 
laboratory purposes can be secured 
as just described. Absolutely pure 
cultures can be secured only by observ¬ 
ing all the precautions necessary in 
bacteriological work. 

Achlya is similar and equally good 
for illustrative purposes. It is found 
on insects, fishes, dead fish eggs, and 
on algae. The zoospores escape in a 
mass, which, for a short time, is held 
together by a transparent pellicle; in 
Saprolegnia the zoospores swarm sepa¬ 
rately. In Saprolegnia , the new spor¬ 
angia grow up through the empty ones; 
in Achlya , the later sporangia arise on lateral branches below the 
earlier ones. Dictyuchus and Olpidium often appear when one is 



Fig. 54. — Saprolegnia: A, sporan¬ 
gium; in B and C the upper cell is an 
oogonium; the cells below are probably 
sporangia. The material was grown 
on flies by Miss Mary Norton at mid¬ 
dle Granville, New York, and was fixed 
in formalin acetic acid and stained in 
Magdala red and anilin blue. X150. 











‘214 


METHODS IN PLANT HISTOLOGY 


trying to get Saprolegnia or Achlya. The fixing and staining de¬ 
scribed for Saprolegnia will give good results with the other genera. 

Albugo.—This fungus is quite common on Cruciferae, where the 
white “blisters” or “white rust,” Albugo Candida, form quite con¬ 
spicuous patches. Affected portions of leaves and stems should be 
fixed in chromo-acetic acid and cut in paraffin. Sections 5 m or less 



Fig. 55.— Albugo Candida: A, small portion of a section of a blister showing coenocytic 
mycelium, conidiophores, and multinucleate conidia; B, a young oogonium with large coenocentrum 
and many nuclei: C, later stage, after differentiation into a central ooplasm, surrounded by the multi¬ 
nucleate periplasm; all of the nuclei of the ooplasm, except one, have disorganized. Fixed in the 
special chromo-acetic acid solution and stained in iron-alum haematoxylin and orange. X780. 

in thickness will be found most satisfactory. Stain in iron-alum 
and counter-stain lightly with orange (Fig. 55). 

The white blisters cause little distortion, but are easily recognized 
by their color; the oogonia do not cause any change in color, but they 
cause great distortion in the pods or stems, so that these organs may 
reach several times their normal size. Parts only slightly distorted 
should be selected, as well as the extreme cases; otherwise, you will 



FUNGI 


215 


secure only old fertilized eggs, with very few of the younger stages. 
The oosporic phase of Albugo bliti is easily recognized on Amaranthus, 
especially on A. retroflexus , where the oospores may be seen with the 
naked eye by holding the leaf up to the light. The oospores usually 
occur in more or less circular patches upon the leaf. When they 
occur among the floral structures, there is often a slight reddish colora¬ 
tion. Unfortunately for the collector, it is very seldom that any 
red coloration in Amaranthus is due to the desired material. 

The oosporic stage of Albugo Ipomeae , on the morning-glory, 
causes extreme distortion of the stem. For sections, it is wed to 
cut out small pieces of the cortex, rather than to fix larger pieces of 
the stem. 

HEMIASCOMYCETES 

Saccharomyces.—Formerly it was considered rather difficult to 
demonstrate the nucleus of the yeast cell. With fresh-growing yeast 
the following method by Wager made the classical demonstration. 
Fix in a saturated aqueous solution of corrosive sublimate for at 
least 12 hours. Wash successively in water, 30 per cent alcohol, 
70 per cent alcohol, and methyl alcohol. Place a few drops of alcohol 
containing the cells on a cover, and when nearly dry add a drop of 
water. After the yeast cells settle, drain off the water and allow 
the cells to dry up completely. Place the cover, or slide, with its 
layer of cells in water for a few seconds, and then stain with a mixture 
of fuchsin and methyl green, or fuchsin and methylin blue. Mount 
in glycerin or in balsam. 

With modern methods, there should be no more difficulty than 
in demonstrating the nucleus in the Cyanophyceae or in the mycelium 
of the Phycornycetes. Use an abundance of vigorously budding 
material, so that you can afford to lose most of it and still have a 
plenty left: Fix in the special chromo-acetic-osmic-acid solution and 
stain in iron-alum haematoxylin. Use the Venetian turpentine 
method, or imbed in paraffin and cut sections about 5 p. 

To obtain the spore stage, put a cake of Fleischmann’s yeast in 
a mixture of equal parts of grape juice and distilled water; add 1 g. 
of peptone and allow to bud freely over night at 30° C.; place the 
material in a plaster-of-Paris cup with a depression, and put the 
cup in a small Stender dish with water coming nearly to the top 
of the cup. In 60 to 70 hours there should be abundant spore 
formation. 


216 


METHODS IN PLANT HISTOLOGY 


ASCOMYCETES 

This group, popularly known as the “sac fungi,” contains an 
immense number of saprophytic and parasitic forms. The green 
mold on cheese and leather, the leaf curl of peach, the black knot of 
cherry and plum, and the powdery mildews are familiar to everyone. 
The few objects selected will enable the student to experiment, but he 
must not be discouraged if success does not 
crown the first attempt, for some members 
of the group present real difficulties. 

Peziza.—The Pezizas and related forms 
are fleshy, and present but little difficulty 
in fixing, cutting, or staining. They are 
abundant in moist places, on decaying 
wood, or on the ground. The apothecia 
have the form of little cups, which are 
sometimes black and sometimes flesh- 
colored, but often orange, red, or green. 

For general morphological work it is 
better to tease out fresh or preserved 
material. Such views as that shown in 
Figure 56 are easily obtained in this way. 
For permanent preparations showing such 
views, it is better to stain in bulk in alum 
carmine or in Delafield’s haematoxylin, 
and then tease out the asci in glycerin 
or balsam. Sections showing the entire 
ascus should be 10 to 15 n, in thickness. 

For the free nuclear division in the 
ascus, and also for the development of the 
ascospores, Flemming’s weaker solution, 
followed by the safranin, gentian-violet, 
orange combination has given excellent results. The centrosomes, 
especially at the first division in the ascus, are sharply defined and 
the radiations are conspicuous. With iron-alum haematoxylin and 
orange, the nuclear detail and the centrosomes are better, but the 
spindle and radiations are not so sharply defined. For such details, 
sections should not be thicker than 5 and 3 m will give a clearer view. 

Morchella esculenta is very good for the development of the ascus 
because the nuclei are very large. 



Fig. 56 .—Peziza odorata: three 
asci and many paraphyses; fixed 
in corrosive sublimate, stained in 
bulk in alum carmine, teased out, 
and mounted in balsam. X245. 
















FUNGI 


217 


For showing the ascogonium, ascogenous hyphae, and the origin 
of the asci, nothing is better than Pyronema. Fix in formalin, acetic 
acid (10 c.c. formalin, 5 c.c. acetic acid, and 85 c.c. water) for 24 
hours or more; wash in water and stain in eosin. Or, fix in the special 
chromo-acetic-osmic solution and stain in iron-alum haematoxylin. 
In either case, use the Venetian turpentine method and tease the 
material so as to obtain instructive views. 

Eurotium. —Eurotium with its conidial stage, Aspergillus , is a 
very common mold found on bread, cheese, decayed and preserved 
fruit, etc. In the conidial stage it is green and in the ascosporic 
stage yellow, reddish yellow, or reddish brown. Aspergillus is 



Fig. 57. — Aspergillus: from material growing on a hectograph pad; fixed in chromo-acetic 
acid, stained in eosin, and mounted in glycerin; A-E, successive stages in development. X375. 
All such material is more satisfactory when mounted in Venetian turpentine. 

almost sure to appear upon bread which is kept moderately moist, 
because the conidia are usually abundant in the atmosphere. If 
the bread be wet with a 10 per cent solution of cane-sugar or with 
grape juice, this stage appears sooner and in greater abundance. 
A temperature of 22° to 30° C. is also a favorable condition. 

The perithecial stage is not found so frequently, but can sometimes 
be secured by examining moldy preserves. The sexual stage has been 
induced. Soak a piece of bread in a 20 per cent solution of grape-sugar 
in grape juice; upon this sow the spores and keep at a temperature of 
about 28° C. After 4 or 5 days, begin to examine. A 40 per cent solution 
of cane-sugar in the juice of prunes is also a good nutrient solution. 

For class use or for permanent preparations it is best to select 
rather young material which shows various stages in development, from 
the swollen end of the hypha to the ripe spore (Fig. 57). Permanent 
preparations of the conidial stage, as shown in Figure 57, and also of the 
coiled twisted filaments which initiate the ascosporic stage, should be 
made by the Venetian turpentine method or by the glycerin method. 





218 


METHODS IN PLANT HISTOLOGY 


Fix in 1 per cent chromo-acetic acid (1 g. chromic acid and 1 c.c. 
acetic acid and 100 c.c. water) for 24 hours; wash in water 24 hours, 
stain sharply in eosin, transfer to 10 per cent glycerin, and follow 
the Venetian turpentine method. 

Material may be fixed in corrosive sublimate-acetic acid (cor¬ 
rosive sublimate 2 g., glacial acetic acid 2 c.c., and water 100). Use 
it hot (85° C.). One minute is long enough. Wash in water and 
add, a few drops at a time, the iodine solution used in testing for 
starch. At first, the brownish color caused by the iodine will dis¬ 
appear, but after a certain amount has been added the brownish 
color will remain. Stain in eosin or iron-haematoxylin and follow 
the Venetian turpentine method. 

A very rapid method for this and for similar small filamentous 
forms may be added. Forms as large as Thamnidium elegans can 
be mounted successfully by this method. 

1. 100 per cent alcohol, 2 minutes. 

2. Eosin (aqueous), 2 minutes. 

3. 1 per cent acetic acid, 2 to 10 seconds. 

4. Mount directly in 50 per cent glycerin and seal. 

If the material gets through the first three stages without shrinking 
but collapses at the fourth, put it into 10 per cent glycerin and allow 
it to thicken, following the Venetian turpentine method. 

The earlier perithecial stages are more instructive when mounted 
whole; but later stages, even before the formation of the asci, are 
very unsatisfactory by this method, and should be cut in paraffin. 

Penicillium.—This green mold is found everywhere upon de¬ 
caying fruit, upon bread, and upon almost any decaying organic 
substance. Material is even more easily secured than in case of 
Aspergillus , and Penicillium is an easier type for laboratory study. 
Such a satisfactory study can be made from the living material that 
it is hardly worth while to fix and stain. The very rapid method 
described for Aspergillus will furnish good mounts if permanent 
preparations are desired. 

The Erysipheae.—The mildews are found throughout the summer 
and autumn on the leaves of various plants. Some of the most 
abundant forms are Microsphaera alni on the common lilac; Sphaero- 
theca castagnei on Bidens frondosa and other species, on Erechtites 
hieracifolia , and on Taraxacum officinale; Uncinula necator on 
Ampelopsis quinquefolia, and U. salicis on Salix and Populus; 


FUNGI 


219 


Erysiphe commune on Polygonum aviculare; and Erysiphe cichoria- 
cearum on numerous Compositae and Verbenaceae. Podosphaera 
may be found on the leaves of young cherry trees and apple trees, 
and on young shoots of older trees. The infected leaves are likely 
to be more or less deformed. Phyllactinia is sometimes abundant on 
leaves of Alnus incana. It is also found on Celastrus, Desmodium, 
Typha, and on various members of the Amentiferae. For herbarium 
purposes they may be preserved by simply drying the leaves under 



Fig. 58.— Uncinula necator on Ampelopsis quinquefolia: A, four asci containing ascospores 
have been forced out by pressing on the cover; stained in fuchsin and mounted in balsam; B, a 
conidiospore; and C, an appendage of Microsphaera alni, drawn from living material. X192. 

light pressure. When needed for examination the leaf should be 
soaked in water for a few minutes, after which the perithecia may 
be scraped off and mounted in water. In mounting great care must 
be taken not to break off the appendages. The asci may be forced 
out by tapping smartly on the cover (Fig. 58). 

For permanent mounts of entire perithecia with appendages, fix 
in 5 per cent formalin 24 hours, wash in water 1 hour, stain in aqueous 
eosin 24 hours, remembering to keep all solutions slightly acid. 
Use the Venetian turpentine method. If chromic acid, corrosive 









220 


METHODS IN PLANT HISTOLOGY 


sublimate, or alcohol be used for fixing, the appendages become brittle 
and very easily break off. However, the chromo-acetic mixtures are 
better if it is desired to make paraffin sections showing the develop¬ 
ing of the perithecium with its asci and spores. For this purpose 
the omnipresent Erysiphe commune on Polygonum aviculare is excep¬ 
tionally favorable, because, after the material has been fixed and has 
been brought into alcohol, the whole mycelium, with the develop¬ 
ing perithecia, may be stripped from the leaf without the slightest 

difficulty, thus avoiding the 
necessity of cutting the leaf 
n in order to get the fungus. 
)l Material in which the peri- 
I thecia are still white or yel¬ 
lowish contain stages up to 
k the formation of the uninu¬ 
cleate ascus; brownish peri- 
I thecia show the development 

of ascospores, and dark- 
brown or black perithecia 
contain the mature asci with 
fully developed ascospores. 
In early stages while the 
perithecia are still yellow or 
a very slightly brownish, the 
C1 material can be stripped off 
from the leaves before fixing. 
An air-pump will remove any air. Use iron-alum haematoxylin and 
orange, or the safranin, gentian-violet, orange combination. Sections 
thicker than 5 y will be hard to stain effectively. 

The Xylariaceae.—Most of these forms, in their mature con¬ 
dition, are black. In younger stages the color is lighter, often show¬ 
ing gray, brick-red, or brownish tints. Nummularia is common on 
dead branches of beech, elm, oak, locust, and other trees. It is gen¬ 
erally flat, orbicular, or elliptical in form. Ustilina is a crustaceous 
form, rather diffuse and irregular in shape. It is most common 
on the roots of rotten stumps. Hypoxylon is more or less globose 
in form, and the color is brick-red, brown, or black. It is found on 
dead twigs and bark of various trees, especially beech, and is more 
abundant in moist situations. Xylaria (Fig. 59) is found on decaying 









FUNGI 


221 


stumps and logs, and often apparently on the ground, but really 
growing on twigs, wood, and bark just under the surface. When 
mature it is black outside and white or light-colored within. When 
young, it is easily cut in paraffin; in some forms the ascospores are 
fully formed before the stroma becomes hard enough to occasion any 
difficulty in cutting. When the stroma becomes black, many mem¬ 
bers of the Xylariaceae become very hard and brittle, so that sections 
are likely to be unsatisfactory. For general morphological study 
it is better to break the stroma transversely and examine with the 
naked eye and with a pocket lens. The asci with their spores can be 
teased out and mounted in water. For permanent preparations, soak 
the stroma for a month in equal parts of 95 per cent alcohol and 
glycerin; then cut sections, and, after leaving them in glycerin for 
a day or two, mount in glycerin jelly. It is better not to stain the 
old stages. For illustrative purposes, select forms which can be 
cut in paraffin. The method just given merely shows that such 
material can be cut. 

LICHENS 

The lichens are usually regarded as difficult forms. In younger 
stages they occasion no trouble, but an old apothecium or a leathery 
thallus often fails to cut well. By employing the gradual processes 
already described in chapter ix, satisfactory sections should be 
obtained from thalli and mature apothecia of Physcia, Usnea, Sticta, 
Parmelia, and Peltigera. Collema and other lichens of such gelatinous 
consistency, while they cut readily, show a strong tendency to wrinkle. 

Cyanin and erythrosin is a very good stain for lichens. The 
algae stain blue and the filaments of the fungus take the red. Where 
the association of the alga and the fungus is rather loose, as in 
Dichonema, more satisfactory mounts can be made by staining in 
eosin, or haem-alum and eosin, and then teasing slightly with needles 
and mounting in glycerin. 

BASIDIOMYCETES 

This is an immense group, of which the smuts, rusts, mush¬ 
rooms, toadstools, puffballs, and bracket fungi are the most widely 
known representatives. 

The Smuts (Ustilagineae).—The smuts are abundant on wheat, 
oats, corn, and various other plants. 


222 


METHODS IN PLANT HISTOLOGY 


The smuts may be studied in the living material. The following 
method, described by Ellis, is worth remembering: A supply of 
smutted barley may be obtained by sowing soaked, skinned barley 
that has been plentifully covered by Ustilago spores. In such 
material it is easy to trace stages in the development of spores. 
Freehand sections of ears about 12 mm. long show the mycelium and 
spore clusters. If smutted ears be removed and kept floating on the 
water, the spores continue to develop and often germinate. For 
paraffin sections desirable stages should be fixed in Flemming’s fluid 
or picro-acetic acid. Delafield’s haematoxylin, followed by a very 
light touch of erythrosin or acid fuchsin, will give a good stain. 

For a study of the germinating spores and conidia, cultures may 
be made in beerwort on the slide or in watch crystals. Harper’s 
method of making preparations from such material is ingenious 
and is valuable in making mounts of various small plant and animal 
forms. A drop of the material is taken up with a capillary tube and 
is then gently blown out into a drop of Flemming’s weaker solution 
(15 minutes to 1 hour was sufficient for the fungus spores). Cover a 
slide with albumen fixative, as if for sections. A drop of the material, 
without previous washing, is drawn up into the capillary tube and 
touched lightly and quickly to the surface of the albumen. A series of 
such drops, almost as small as the stippled dots in a drawing, may 
be applied to the slide. The fixing agent may now be allowed to 
evaporate somewhat, but the preparation must not be allowed 
to dry. As the slide is passed rapidly through the alcohols, the 
albumen is coagulated, and the preparation may be treated just as if 
one were dealing with ribbons of sections. 

The Rusts (Uredineae). —Puccinia graminis , the common rust of 
wheat and oats, is familiar to everyone. The uredospores, or summer 
spores, known as the red rust, and the winter spores, known as the 
black rust, are found in unfortunate abundance, but the aecidium 
stage on the barberry is not necessary for the vigorous development 
of rust in the United States, and it is not nearly so prevalent as the 
red- and black-rust stages. When found, it may be so abundant that 
most of the leaves of the barberry are spotted with the cluster cups. 
It is a curious fact that wheat and oats may be quite free from the 
red and black rust in localities where the aecidium stage is very 
abundant; and that the rust stages may be most destructive where 
there are no barberry bushes. But no one doubts that the aecidium 


FUNGI 


223 


stage on barberry belongs in the life-history. The aecidium on bar¬ 
berry cuts easily in paraffin (Fig. 60). If the aecidium stage is not 
easily available, there are various aecidia which are just as good, or 
even better, for morphological study. The aecidia growing on 
Euphorbia maculata (spotted spurge) are abundant and are very easy 
to fix and cut. The infected plants are also very easily recognized, 
normal plants having the prostrate habit, while infected plants 
become erect and the internodes become greatly elongated. Aecidia 
growing on Arisaema triphyllum (Jack-in-the-puplit) are also easy 



Fig. 60 . —Puccinia graminis: photomicrograph of aecidium stage on barberry. Fixed in 
chromo-acetic acid and stained in cyanin and erythrosin: Eastman Commercial Ortho film, Wratten 
E filter (orange); arc light; Spencer 16-mm. objective N.A. .25; Bausch and Lomb projection 
eyepiece; exposure, £ second. X47. Negative by Dr. P. J. Sedgwick. 

to cut. The Aecidium on Hepatica has large nuclei and affords 
particularly good views of the intercalary cells. 

The special chromo-acetic-osmic-acid solution is recommended 
for fixing and iron-haematoxylin with a faint touch of orange is a 
satisfactory stain (Fig. 61). 

It is rather difficult to get good sections of uredospores and teleu- 
tospores of Puccinia graminis , because the leaves of wheat and oats 
are refractory objects to cut (Fig. 62). For illustrative purposes, 
soak the leaves, scrape off the spores, and study without sectioning. 
For sections, select species growing on less refractory hosts. 

Everyone who studies the rusts should attempt to germinate the 
uredospores and teleutospores. For this purpose the hanging-drop 





224 


METHODS IN PLANT HISTOLOGY 


culture may be employed, as described in the chapter on temporary 
mounts (chap. v). The uredospores germinate readily all summer, 
but in most forms teleutospores will germinate only in the spring 
following their maturity. However, the teleutospores of “lepto” 
species, like Puccinia xanthii on Xanthium, canadense (cocklebur), 


will germinate as soon as they ripen, and will serve equally well for 
study. If a particularly good specimen 
is secured, it may be preserved by the 
method previously described for desmids, 
except that in this case it might be worth 
while to attempt staining with Mayer’s 
haem-alum or with eosin. 

Gymnosporangium, which is rather 
common on Juniperus virginiana (red 
cedar), forms its basidia in the “cedar- 
apple” stage. Bring the cedar apples into 
the laboratory in late winter or early spring 
and put some into a dish of water. The 
yellowish, gelatinous strands with the 
germinating teleutospores may appear 
within 24 hours. The various stages are 
easily recognized under a low-power dry 
lens without even crushing the gelatinous 
masses. Fix in 10 per cent formalin for 
24 to 48 hours. The material may be left 
here indefinitely. In chromo-acetic acid 
the gelatinous substance goes to pieces. 
Stain in iron-alum haematoxylin and 
follow the Venetian turpentine method; 
or, when the thick glycerin is reached, 
mount in glycerin jelly. The material 
spreads out better and fewer basidia are torn off from the teleutospores 
in glycerin jelly than in the Venetian turpentine. 

The interesting nuclear conditions in the life-history of a rust 
which has uredospores, teleutospores, and aecidiospores are not diffi¬ 
cult to demonstrate. After the uredospore or teleutospore stage has 
been fixed in chromo-acetic acid and washed in water for an hour, 
treat with 10 per cent hydrofluoric acid in water for an hour and then 
continue the washing in water for 24 hours. If fixed in alcoholic 



Fig. 61 .—Aecidium on Hepatica: 
fixed in chromo-acetic acid with a 
little osmic acid, and stained in safra- 
nin, gentian-violet, orange; from a 
preparation by Dr. Wanda Pfeiffer 
Vestal. X950. 






FUNGI 


225 


corrosive sublimate and acetic acid, rinse in 50 per cent alcohol, 
then treat with 10 per cent hydrofluoric acid in 50 per cent alcohol, 
wash in two or three changes of 50 per cent alcohol. In either case, 
imbed in paraffin. Cut about 5 n in thickness and stain in iron-alum 
haematoxylin. A rather strong stain of orange in clove oil will 
make it easier to trace the mycelium in the host. Uredospores and 
teleutospores, both young and old, are shown in Figure 62. 

Various stages in the germination of the teleutospores of Gymno- 
sporangium are shown in Figure 63. Reduction of chromosomes takes 
place at the first two divisions of the nucleus in the basidium. The 
sterigmata in this species are very large. 

The Fleshy Fungi.—-For habit study, nothing is equal to fresh 
material; for second choice, buy canned “mushrooms” (usually 
Agaricus campestris) at the grocery; forms not readily available in 
field or grocery may be preserved in formalin alcohol (6 c. c. of formalin 
to 100 c.c. of 50 per cent alcohol). When formalin is used in water, 
the fungi become too soft. Larger forms of the mushroom, puff¬ 
ball, and bracket types may be dried in an oven. The circulation of 
air should be good and the temperature should be kept at about 50° C. 
After drying, the fungi should be poisoned. 

For sections, Gilson’s fluid deserves more recognition than it has 
received. It is particularly good for soft forms, like Tremella. 


Gilson’s Fluid.— 

95 per cent alcohol. 42 c.c. 

Water. 60 c.c. 

Glacial acetic acid. 18 c.c. 

Concentrated nitric acid. 2 c.c. 


Corrosive sublimate (saturated solution in water).. 11 c.c. 

Fix about 24 hours and wash in 60 or 70 per cent alcohol. 

Coprinus micaceus is particularly good for a study of gills, basidia, 
and the formation of basidiospores, because it is so small that a single 
section may show a fine series of stages. Gills which are becoming 
brownish at the tip, but which are still white toward the top of the 
cap, will show a splendid series of stages. For fixing, cut out pieces 
of the gills 1 cm. long and 3 mm. thick. Such material fixes well in 
Flemming’s weaker solution. Cut paraffin sections perpendicu¬ 
lar to the gills. To show the four basidiospores, sections should 






226 


METHODS IN PLANT HISTOLOGY 



Fig. 62.— Puccinia, graminis: A, uredospores on oats, showing binucleate mycelium and 
binucleate uredospores with binucleate stalk cells; c, chloroplasts, and n, nuclei of host cells; 
B, C, and D, stages in the development of the teleutospore; the two nuclei of the binucleate cells 
in C fuse to form the uninucleate cells of D. X780. 



Fig. 63.— Gymnosporangium: A, beginning of germination of teleutospore; B, later stages, 
the lower basidium showing the nucleus in the synapsis stage of the heterotypic mitosis, and the 
upper basidium showing the four-cell stage following the homotypic mitosis; C, the two figures of 
the homotypic mitosis; D, E, F, and G, later stages showing the formation of basidiospores, many 
of the basidiospores being binucleate— g, gelatinous stalk; b, basidiospore; s, sterigma. Fixed 
in 10 per cent formalin and stained in iron-alum haematoxylin. X800. 















FUNGI 


227 


be 10 to 15/x thick; to show details of nuclei, 3 /z is thick enough 
(Fig. 64). 

In Hydnum and Polyporus , cut out pieces about 3 or 4 spines or 
3 or 4 pores in width and about 1 cm. long. A rectangular piece 
which will allow the transverse 
sections of the spines or pores to 
be about 4 mm. wide and 1 cm. 
long cuts better than a piece which 
will give square sections. 

In Boletus , simply strip off 
the hymenium and cut into pieces 
which will give transverse sections 
of the tubes. 

In Lycoperdon, Bovista , 

Geaster , and Scleroderma , longi¬ 
tudinal sections of the entire 
fructification can be cut in paraf¬ 
fin as long as the fresh material is 
easily sliced with a Gillette blade. 

Young stages of Cyathus, Cru- 
cibulum, and Nidularia cut easily 
in paraffin; somewhat older stages 
can be cut in celloidin, but mature 
stages fail to cut by any of our 
present methods. 

It is often desirable to secure 
differential staining of the fungus 
and its host. Some of the methods previously mentioned secure 
this result and give excellent detail, but do not make the mycelium 
stand out sharply in contrast with the tissues of the host. A special 
method by B. T. Dickson generally gives a good differentiation. He 
uses Magdala red and light green, using the Magdala red in a 2 per 
cent solution in 85 per cent alcohol, and the light green in a 2 per cent 
solution in clove oil to which has been added a few drops of abso¬ 
lute alcohol. His schedule is as follows: 



Fig. 64.— Coprinus: young basidia with 
four nuclei which, later, pass into the spores; 
fixed in chromo-acetic acid and stained in 
safranin, gentian-violet, orange. X780. 


1. Dissolve paraffin in xylol and wash in absolute alcohol. 

2. Wash in 95 and 85 per cent alcohol. 

3. Stain with Magdala red 5 to 10 minutes. 

4. Remove surplus stain and wash in 95 per cent alcohol. 









228 


METHODS IN PLANT HISTOLOGY 


5. Stain in light green in clove oil for 1 to 3 minutes. 

6. Wash in absolute alcohol, or in carbol turpentine. 

7. Clear in xylol and mount in balsam. 

The time factors will vary slightly with different material. If 
the light green overstains slightly it may not interfere much with the 
differentiation. Mycelium, spores, amoebae, and bacterioidal tissue 
stain red and the host tissues green. If tissues do not stain readily, 
mordant in a 1 per cent solution of potassium permanganate in water 
for 2 to 5 minutes, wash in water, pass through the alcohols to 85 per 
cent alcohol, and stain. The mordant does not keep. 

This combination has given good results with the following ma¬ 
terial: Plasmodiophora brassicae, legume tubercles , Albugo Candida , 
Phytophora infestans , Uromyces caryophyllinus, Puccinia graminis , 
and many others. 


CHAPTER XIX 


BRYOPHYTES 


The Bryophytes, comprising the two groups of Liverworts 
(.Hepaticae ) and Mosses (Musci), present a great diversity of struc¬ 
ture, some being so delicate that good preparations are very uncer¬ 
tain, while others are so hard that it is difficult to get satisfactory 
sections. Between these extremes, however, there are many forms 
which readily yield beautiful and instructive preparations. 

If but one fixing agent should be suggested for the entire group, 
it would be chromo-acetic acid with 1 g. chromic acid and 2 c.c. 
acetic acid to 100 c.c. of water. It should be allowed to act for at least 
24 hours. For morphological study, Dr. Land uses a formalin 
alcohol solution (6 c.c. commercial formalin to 100 c.c. of 50 per cent 
alcohol), which he has tested in extensive collections in various 
tropical regions, where it has been impracticable to use the chromic 
series, with its tedious washing and changing of alcohols. Material 
may be left in the formalin alcohol solution until needed for use, a 
convenience which will hardly be appreciated by those who are 
always within reach of a laboratory. 

For general study, the small, delicate forms may be mounted 
whole in Venetian turpentine. 

Instead of treating forms in a taxonomic sequence, we shall 
consider first the gametophyte structures under the headings thallus, 
antheridia, and archegonia, and shall then turn our attention to the 


sporophyte. 


HEPATICAE 


Some of the liverworts are floating aquatics, but most of them 
grow on logs or rocks or upon damp ground. They are found at their 
best in damp, shady places. Many of them may be kept indefinitely 
in the greenhouse. Riccia, Marchantia, Conocephalus, Asterella, and 
many others vegetate luxuriously, and often fruit if kept on moist 
soil in a shady part of the greenhouse, and they do fairly well in the 
ordinary laboratory if covered with glass and protected from too 
intense light. Riccia natans is a valuable type for illustrative 

229 


230 


METHODS IN PLANT HISTOLOGY 


purposes. It floats freely on the surfaces of ponds and ditches. 
Early in the spring (during April in the Chicago region) it produces 
antheridia; then, for a short time (about the first of May) both 
antheridia and archegonia, and still later only archegonia. Sporo- 
phytes then appear as black dots along the grooves. After the 
spores are shed, the thallus remains sterile for the rest of the season. 
Marchantia and similar forms are not difficult to establish out of 
doors. A rather damp, shady spot close to the north side of a building 
is best. Scrapings from a board which has been nearly burned 
up make the best fertilizer to scatter on the soil, if one is to cultivate 
Marchantia. Such freezing as Marchantia receives in the vicinity 
of Chicago does not prevent it from appearing again the next spring. 
If it is desirable to have material throughout the year, the out-of-door 
culture may be made in a box which can be brought into the labora¬ 
tory or greenhouse in the winter. A box 3 feet long, 2 feet wide, 
and 1 foot deep will be convenient. It should have a glass cover; 
an old window will do. There should be about 6 inches of dirt in 
the box. A mixture of sand, loam, and charred scrapings will make 
a good substratum for Marchantia. If one is to raise liverworts in 
the laboratory, it is absolutely necessary to note carefully the condi¬ 
tions under which they grow in the field. 

The living plants are very desirable, since they not only furnish 
the best possible material for habit work and the coarser microscopic 
study, but they also enable one to secure complete series in the 
development of the various organs. 

If even a small room in a greenhouse is available, liverworts can 
be grown in great variety and abundance. On one side of the room, 
have a pile of rocks. Half of this space should be occupied by lime¬ 
stone rocks, held in place with as little mortar as possible. There 
should be some shale and some porous red brick. The whole should 
be arranged so that water may trickle down from above. A pipe 
with holes xV inch in diameter will furnish enough water. The other 
three sides may be built up of various rocks, and some clay, so as to 
form a table about 1 m. high. A small fountain, with a bowl a 
couple of feet in diameter, built of rocks, will add to the efficiency. 
If a few well-supported cement tanks be placed above the principal 
pile of rocks, Isoetes and all the water ferns may be grown there, 
besides Elodea, Myriophyllum, Chara, and other forms constantly 
needed in laboratory work. 


BRYOPHYTES—HEPATICAE 


231 




B 


The Thallus.— In many cases it will not be necessary to make a 
special preparation for the study of the thallus, since preparations of 
antheridia, archegonia, or sporophytes may include good sections of 
vegetative portions. This is particularly true of forms like Riccia, 
where the various organs are not raised above the thallus. In forms 
like Marchantidj where the antheridia, archegonia, and sporophytes 
are borne upon stalked receptacles, it is better to make separate 
preparations to show the structure of the mature thallus. Sections 
intended to show the structure of the mature thallus should be 15 to 
25 fjL in thickness, but sections to show the growing point and develop¬ 
ment of the thallus 
should not be thicker 
than 10 fjL. The apical 
region of the Junger- 
manniaceae (Figs. 65, 
66) affords an excellent 
opportunity for study¬ 
ing the development of 
the plant body from a 
single apical cell. If 
mixtures containing 
osmic acid are used for 
fixing, there may be 

difficulty in the staining, even after using peroxide of hydrogen. 

Chromo-acetic mixtures, without osmic acid, are better for the 
apical region. Chromo-acetic acid, followed by Delafield’s haema- 
toxylin, is good for the apical cells and developing regions, but a 
light counter-stain with erythrosin improves preparations of the 
mature thallus. Safranin, with light green, or safranin with crystal- 
violet, will give clear views of the growing region. The latter combi¬ 
nation is particularly good for forms like Pellia , where even the apical 
cell is more or less vacuolated, since it not only brings out the cell 
walls, but stains plastids and other cell contents (Fig. 66). The 
chloroplasts and leucoplasts are well differentiated by this stain. 
After corrosive sublimate-acetic, a vigorous staining in a mixture of 
acid fuchsin and iodine green often brings out the walls very sharply. 
After corrosive sublimate-acetic the material may be stained in bulk 
with alum cochineal or alum carmine, thus giving fairly good prepara¬ 
tions and saving considerable labor. 


Fig. 65. —Ptilidium ciliare: A, longitudinal section, 
and B, transverse section of the apical region of the leafy 
gametophyte. Fixed in chromo-acetic acid and stained in 
Delafield’s haematoxylin. X420. 



232 


METHODS IN PLANT HISTOLOGY 


Antheridia.—It is not difficult to get good preparations showing 
the development of antheridia. In forms like Conocephalus, Asterella, 
Pellia , etc., cut out small portions of the thallus bearing the anther¬ 
idia. The piece should not be more than 1 cm. long and 5 mm. 
wide, preferably smaller. For the development of the antheridia of 
Marchantia, select young antheridiophores which still lie close to the 
thallus. With the antheridiophore, cut out a small piece of the 
thallus, about 5 mm. in length. For general development, cut 10 p, 
but for details of spermatogenesis, sections should not be thicker 



Fig. 66. —Pellia epiphylla: photomicrograph of apex of gametophyte showing apical cell 
and segments; fixed in chromo-acetic-osmic acid and stained in safranin, gentian-violet, orange. 
The negative was made by Dr. Kohler at the Zeiss factory in Jena, Germany. 


than 5 p (Fig. 67). Sections should be stained in iron-alum haema- 
toxylin. The cells are very small and the contents very dense; 
consequently, the staining must be very critical to show the blepharo- 
plasts and chromosomes or, in later stages, the transformation of 
spermatids into sperms. If the material is perfectly infiltrated and 
imbedded, a constant cooling with ice should make it possible to 
secure smooth sections as thin as 3 p, or even 2 p. 

If sperms are found escaping, transfer them to a small drop of 
water on a clean slide, invert the drop over a 1 per cent solution 
of osmic acid for 2 or 3 minutes, allow the drop to dry up, pass the 
slide through the flame 2 or 3 times, as in mounting bacteria, and 





BRYOPHYTES—HEPATICAE 


233 


then stain sharply in acid fuchsin. This should show the general 
form of the antherozoid, and will usually bring out the cilia. 

The Archegonia. The methods for archegonia are practically the 
same as for antheridia. Too much stress cannot be laid upon the 
importance of carefully selecting the material. Use very small 
pieces, and, before placing them in the fixing agent, trim them to 
such a shape that the position of the archegonia will be known 
accurately even after the pieces are imbedded in paraffin. Since air 
is likely to be caught between the perigynium and the archegonium, 



Fig. 67 .—Marchantia polymorpha: early stages in the development of antheridia; from an 
unpublished drawing by Dr. W. J. G. Land. X600. 

it is worth while to use an air-pump as soon as the material is put 
into a chromo-acetic mixture. With the formalin alcohol solution, 
material is likely to sink promptly and the pump is not necessary. 

For stages like those shown in Figure 68A and B, safranin with 
anilin blue or light green is a good stain and 7 to 10 n is about the 
right thickness. 

For stages .like C, in such forms as Marchantia, where the necks 
are long and often somewhat curved, it is better for general purposes 
to use sections about 15 ju in thickness. If it is desired to obtain 
preparations showing the cutting off of the ventral canal cell, the 





234 


METHODS IN PLANT HISTOLOGY 


development of the oosphere, and the process of fertilization, the 
sections should be from 3 to 6 m in thickness. 

For archegonia containing young embryos, like that shown in D, 
Delafield’s haematoxylin without any counter-stain gives beautiful 
preparations when the staining is well done. It is easier for the 
beginner to get good preparations with the safranin, gentian-violet, 

orange combination. 

In Riccia natans 
{Ricciocarpus natans) 
the direction of the 
axis of the archego- 
nium at every stage 
in the development 
must be known; 
otherwise, there will 
be few good longitu¬ 
dinal sections. 

In forms like 
Porella and Scapania, 
the involucre cover¬ 
ing the archegonia 
is likely to hold a 
bubble of air, which 
will delay or even 
prevent fixing. The 
best plan is to cut 
off the offending leaf 

witn a pair ot slender-pointed scissors. Sometimes the air can be got 
out with an air-pump. 

The Sporophyte.—Sporophytes in early stages of development 
often yield good preparations without very much trouble, but in 
later stages they are frequently difficult to cut on account of the 
secondary thickening of the capsule wall and the stubborn exine of 
the mature spores. Great care must be taken to get Riccia natans 
into paraffin without shrinking, and the same thing may be said of 
other forms which have such loose tissue with large air cavities. 
Formerly, we resorted to celloidin for stages like that shown in 
Figure 69. The gradual processes already described have obviated the 
difficulty, so that the student should be able to get thin paraffin 




Fig. 68. —Marchantia polymorpha: A, three early stages in 
the development of archegonia—Delafield’s haematoxylin; B, 
young archegonium showing two neck canal cells and the central 
cell before the cutting off of the ventral canal cell—fuchsin and 
methyl green; C, mature archegonium immediately before the 
fertilization period—safranin, gentian-violet, orange; D, young 
embryo—Delafield’s haematoxylin. X400. 














BRYOPHYTES—HEPATICAE 


235 


sections as free from distortion as were the old celloidin sections. 
But even with well-fixed material great care must be taken not to let 
the paraffin get too hot. Remember that in most paraffin ovens the 
temperature is different in different parts of the oven. Do not let 
the temperature of the paraffin go above 53° C., and, preferably, not 
above 52° C. In Riccia natans it is even more difficult to get median 
longitudinal sections of the sporophyte than of the archegonium. 

Sections perpendicular to the groove, 
whether longitudinal or transverse, 
are almost sure not to give median 
longitudinal sections of the sporo¬ 
phyte, and these are the sections the 
beginner is sure to cut. Examine 
the material and note very exactly 
the orientation of the sporophyte; 
then, for fixing, cut out sections about 
2 mm. thick, taking these sections in 
such a plane that paraffin sections 
parallel to the thick section will give 
the desired median longitudinal sec¬ 
tions of the sporophyte. 

In forms like Pellia and Aneura, 
it is desirable to show the sporophyte 
still inclosed in the calyptra (Fig. 70). 
For such sections, we should recom¬ 
mend fixing in formalin alcohol. 
Aqueous fixing agents are likely to 
cause trouble on account of air bubbles. For cytological studies, 
the calyptra must be removed and a thin slab should be cut 
from opposite sides of the capsule to facilitate fixing and infiltra¬ 
tion. Chromo-acetic acid, with the addition of a little osmic acid, is 
best for fixing. In Pellia and Conocephalus the spores are very large 
and have a rather thin wall. Both these genera show a peculiar intra- 
sporal development of the gametophyte, i.e., the gametophyte 
develops to a considerable extent before it ruptures the spore wall 
and before it is shed from the capsule (Fig. 71). Mitotic figures 
during the first three divisions in these spores are exceptionally beauti¬ 
ful and are very easy to stain with the safranin, gentian-violet, 
orange combination, the chromosomes taking a very brilliant red, 



Fig. 69 .—Riccia natans: young spor¬ 
ophyte inclosed in the archegonium; 
spore mother-cell stage — Delafield’s 
haematoxylin. All the cells of the spor¬ 
ophyte, except a single peripheral layer 
(dotted in the figure), produce spores. 
Celloidin section 30 n in thickness. X104. 



236 


METHODS IN PLANT HISTOLOGY 



while the asters take the violet. Achromatic structures are very 
prominent during these three divisions, but become less and less 

conspicuous as division progresses and 
before the intrasporal stage is over, the 
radiations are scarcely demonstrable. 

For the older sporophytes of Mar- 
chantia, it is better not to cut the 
whole receptacle. Remove the radiating 
branches. The sporophytes are in radi¬ 
ating rows, alternating with the branches. 
A piece 2 mm. wide can be cut so as to 
include two of the radiating rows, one on 
each side of the stalk, and such a piece 
will include early stages in other rows. 
By taking such care, you can get median 
longitudinal sections of nearly all the 


Fig. 70 .—Pellia epiphylla: photo¬ 
micrograph of longitudinal section of 
sporophyte still inclosed in the 
calyptra. The deeply lobed spore 
mother-cells are dividing. Eastman 
Commercial Ortho film, Wratten E 
filter (orange); arc light; J. Swift and 
Son 1-inch lens; exposure, 1 second. 
Negative by Dr. P. J. Sedgwick. 
X30. 



Sporophytes. For class Fig. 71 .—Pellia epiphylla: photomicrograph of spore 
work, 5 to 10 IX is a good germinating while still within the capsule. Fixed in 
n r , r p. chromo-acetic-osmic acid, and stained in safranin, gentian- 

tnickness, but lor figures, violet, orange. Negative by Miss Ethel Thomas. X276. 

especially the reduction 

mitoses in the spore mother-cells, the sections should not be thicker 
than 2 or 3 / 1 . 




BRYOPHYTES—HEPATICAE 


237 


Among the Bryophytes no form affords a better opportunity 
for studying the development of spores than Anthoceros, since a 
single longitudinal section of the sporophyte may show all stages, 
from earliest archesporium to mature spores (Fig. 72). The sporo¬ 
phyte is even more difficult to orient than that of Riccia natans. 
Cut a slice 1 or 2 mm. thick, so as to orient the visible portion of 
the sporophyte, and trust to luck for the orientation of the foot. 
For studies like A and B, chromo-acetic material cut 10/x thick 



Fig. 72.— Anthoceros laevis: A, longitudinal section of lower portion of sporophyte imbedded in 
thegametophyte; X45; B, transverse section of lower portion of sporophyte; X200; C, vegetative 
cell from lower portion of the sporophyte; X560; D, spore mother-cell showing three of the four 
chloroplasts with numerous starch grains; the nucleus in the metaphase of the first division; X560. 


and stained in Delafield’s haematoxylin is very good. The starch 
grains in the chloroplasts take a beautiful violet color with the 
safranin, gentian-violet, orange combination. With so many stages in 
a single section, it will be impossible to stain all of them well. A stain 
which will show the mother-cells and their divisions will be too 
deep for the mature spores, and a stain which shows the spores 
well will be too faint for the mother-cells. It is better to stain some 
preparations for one feature and others for another. It is not worth 
while to steer a median course. 









CHAPTER XX 

BRYOPHYTES 

MUSCI 

In general, the mosses are more conspicuous than the liverworts 
and easier to collect. Many of the most desirable forms fruit only 
in the spring, but something can be found throughout the summer 
and autumn and some, like species of Sphagnum, pass the winter 
with the antheridia and archegonia in advanced stages of development. 

Material is more troublesome to fix than in the liverworts because 
small bubbles of air hinder penetration of the fixing fluid. Use 
an air-pump. Older archegonia and capsules which have turned 
brownish add to the difficulties of the technic. 

The special chromo-acetic-osmic-acid solution, or this solution 
without the osmic acid, fixes well. If an air-pump is not available 
at the time of fixing, Land’s formalin alcohol solution (6 c.c. commer¬ 
cial formalin to 100 c.c. of 70 per cent alcohol) will be more 
satisfactory. 

Protonema. Protonema of some moss can always be found at 
any season. Look for greenish patches resembling Vaucheria. 
Such mats show the developing protonema and young leafy plants. 
Very young mats of moss will also show good protonema, but are 
not likely to show young buds. The brownish bulbils, which are 
quite common in mosses, can be seen with a good pocket lens. The 
little Webern, almost always found on the pots in the fernery or on 
the benches in greenhouses, quite frequently shows this mode of 
reproduction. Protonema is easily grown from spores. 

Permanent mounts are very easily made. Simply wash away 
the dirt with water and put the material into 50 per cent glycerin, 
and let the glycerin concentrate. Mount in glycerin or glycerin 
jelly for permanent mounts. Seal thoroughly. Such mounts, with 
no fixing or staining, may retain the green color for many years. 

If you do not insist upon keeping the green color, much clearer 
mounts can be made by fixing in formalin acetic acid, about 10 c.c. 
of formalin and 5 c.c. of acetic acid to 100 c.c. of water, and staining 

238 


BRYOPHYTES—MUSCI 


239 


in eosin, or in Magdala red and anilin blue. Mount in Venetian 
turpentine. 

Antheridia. -It is easy to find material for a study of antheridia, 
because, in so many cases, the antheridial plants can be detected at 
once without even a 
pocket lens. Funaria, 
with its bunch of anther¬ 
idia as large as a 
pinhead, is extremely 
common everywhere. 

Spring is the best time 
to collect it, but it is 
found fruiting in the 
autumn and sometimes 
in summer; besides, it 
is easily kept in the 
greenhouse, where it 
may fruit at any time. 

Bryum roseum has a 
large cluster of anther¬ 
idia surrounded by ra¬ 
diating leaves, making 
it easy to recognize. 

Other species of Bryum 
and species of Mnium, 
like M. cuspidatum , 
make good sections. 

Polytrichum has a large 
cluster of antheridia 
surrounded by reddish 
leaves, so that the whole 
is sometimes called the 
moss “flower.” In fix¬ 
ing this or the closely 
related Atrichum ( Ca- 
tharinea), cut a small slab from two Sides, so as to leave a flat piece 
to cut for longitudinal sections. This trimming will greatly facilitate 
fixing and infiltration. A single antheridial plant of Poly trichum 
often furnishes a fairly complete series of stages in the development 



Fig. 73 .—Mnium cuspidatum: A, nearly mature antheri- 
dium in the center, with a young archegonium at the right, 
and at the left a still younger stage which may develop into 
either an antheridium or an archegonium; B, a nearly mature 
archegonium with a young archegonium at the right. Fixed 
in formatin, acetic alcohol (formalin 10 c.c., acetic acid 5 c.c., 
50 per cent alcohol, 100 c.c.) and stained in safranin, gentian- 
violet, orange. X240. 




















240 


METHODS IN PLANT HISTOLOGY 


of antheridia. Transverse sections show not only the antheridia, 
but also good views of the peculiar leaf of this genus. In all cases 
the stem should be cut off close up to the antheridia, for many of the 
moss stems, after they have begun to change color, cut like wire. 

Sections to show the development of the antheridium should be 
5 to 10 m in thickness. The safranin, gentian-violet, orange is a good 



Fig. 74. —Funaria hygrometrica: A, apex of young sporophyte showing endothecium and 
amphithecium—chromo-acetic acid and Delafield’s haematoxylin; 10 pt; X420; B, C, and D, 
transverse sections of a sporophyte of the same age as A, taken at different levels; X255.* 


combination (Fig. 73 A). For details of spermatogenesis, sections 
should not be thicker than 3 /jl. Iron-haematoxylin is a better 
stain for the chromatin and blepharoplasts. 

Although sections 20 to 50 /x in thickness can be cut to show 
topography, it is far better to study such stages in the fresh material. 
When a particularly fine view is secured in this way, a permanent 
preparation may be made by putting the piece into 10 per cent glycerin 
without any fixing or staining, and allowing the glycerin to concen¬ 
trate. Then mount in glycerin jelly. 
























BRYOPHYTES—MUSCI 


241 


Archegonia.—Since the necks of the archegonia are usually 
long and more or less curved, it is necessary, for habit work, to cut 
sections as thick as 20 or 30 ju in order to get a view of an arche- 
gonium in a single section (Fig. 735). Mayer’s albumen fixative is 
not likely to held such sections to the slide. Use Land’s fixative. 
Here, as in case of antheridia, it is better to use fresh material, 



Fig. 75 .—Funaria hygrometrica: A, longitudinal section of capsule; B, transverse section of 
capsule of about the same age as A —Delafield’s haematoxylin and erythrosin; 10 /x. The columella, 
archesporium, outer spore case, two layers of chlorophyll-bearing cells, and the beginning of the air 
spaces can be distinguished at this stage. X420. 

putting particularly good pieces into 10 per cent glycerin for glycerin 
jelly mounts. 

For the development of the archegonium, trim away the leaves 
which usually cover the cluster. Fix in chromo-acetic acid with a 
little osmic acid and cut 5 to 10 ^ thick. For a study of the ventral- 
canal cell and fertilization, sections should not be thicker than 5 /jl. 

There is a general impression that the antheridia and archegonia 
of Sphagnum are rare and hard to find. Dr. George Bryan, who made 


























242 


METHODS IN PLANT HISTOLOGY 


an extensive study of Spagnum subsecundum , found that antheridia 
appear in August and archegonia in September. In examining acres 
of this species, he did not find a sterile plant. 1 

Sporophyte.—It is often difficult to get good mounts of sporo- 
phytes. In the younger stages the calyptras are likely to interfere 



Fig. 76—Mnium: upper part of capsule showing two teeth of the peristome cut longitudinally; 
a, annulus. Fixed in formalin alcohol and stained in safranin and light green. X120. 


with cutting, while in the older stages the peristome, or hard wall 
of the capsule, occasions the trouble. If an attempt is made to 
remove the calyptra in young stages, like A of Figure 74, the apex 
of the sporophyte usually comes with it. While picro-acetic acid 
material cuts more easily, chromo-acetic acid followed by Dela- 
field’s haematoxylin gives so much sharper differentiation in stages 

1 Botanical Gazette, 59 :40-56, 1915. 














BRYOPHYTES—MUSCI 


243 


like those shown in Figure 75 that it is better to use ice or Land’s cooler 
and get preparations from chromic material. 

Stages like that shown in Figure 75 are cut with comparative ease, 
for the calyptra is easily removed, and the capsule wall is not yet 
hard enough to occasion any difficulty. Safranin, gentian-violet, 
orange is a good stain. The cell walls stain so sharply that they are 
not obscured by a stain which will bring out the cell contents. 

Later stages, after the peristome 
has begun to differentiate, are likely to 
occasion difficulty in cutting. Bryum 
cuts as easily as any (Fig. 76). For 
the development of the peristome, fix 
in formalin alcohol and stain in safranin 
and anilin blue, or in safranin and 
light green. Safranin and Delafield’s 
haematoxylin is also an excellent stain 
for the older stages in the differentia¬ 
tion of the capsule. 

Beautiful mounts of the peristome 
are easily and quickly made. Take a 
capsule at the stage when the opercu¬ 
lum is just ready to fall off, or has just 
fallen off; with a sharp razor cut off 
the end of the capsule just below the 
line of the annulus; put it into absolute 
alcohol for 10 minutes, clear in clove oil, 
transfer to xylol, and mount in balsam. 

If several are placed on a slide, some 
one side up and some the other, with some complete and some teased 
a little, there will be good views of the entire peristome and also good 
views of teeth and cilia. 

The mature sporophytes of Sphagnum are exceptionally hard to 
cut. It will be worth while to prick the capsule with a needle when 
the material is collected. This will allow the fixing agent to penetrate 
readily, and will also facilitate the infiltration of paraffin or celloidin. 
The puncture causes only a slight damage, and need not reach the really 
valuable portion which is to furnish the median longitudinal sections. 

The younger stages in the sporophyte of Sphagnum like that shown 
in Figure 77, and also the antheridia, archegonia, and the peculiar 
development of the leaves cut easily in paraffin. 



Fig. 77.— Sphagnum: longitudinal sec¬ 
tion of sporophyte showing also the upper 
portion of the pseudopodium and the ca- 
iyptra—Delafield’s haematoxylin. X 24. 





CHAPTER XXI 

PTERID OPHYTES 

This group includes the Lycopodiales, Sphenophyllales, Psilotales, 
Pseudoborniales, Equisetales, and Filicales. The Sphenophyllales 
and Pseudoborniales occur only as fossils and the Psilotales are 
confined to tropical and subtropical regions. The Lycopodiales 
are commonly called club mosses or ground pines, the Equisetales 
are called horsetail rushes or scouring rushes, and the Filicales are 
the common ferns. The Ophioglossaceae, a family of the Filicales, 
is often treated as an order. Two of its genera, Ophioglossum and 
Botrychium, are widely distributed and well known. Material, except 
in the Psilotales, is abundant and so easily recognized that anyone 
who pays a little attention to collecting can, in a single season, get 
a fine supply for a study of the group. Some desirable forms may not 
be present in all localities, but these will be few, and can be secured 
at a reasonable price from those who make a business of collecting. 

The technic for Sphenophyllales will be found under “ Special 
Methods” (chap. xii). Nothing but impressions has yet been found 
in Pseudoborniales. Gametophytes of Psilotales have been found 
only recently. Their young sporangia cut easily, but older stages are 
very refractory and should receive extreme care in dehydrating, 
clearing, and infiltration. No further directions will be given for 
these rather inaccessible orders. 

LYCOPODIALES 

Lycopodium.—The genus is evergreen, and consequently some 
stage in development can be secured at any season. In general, the 
tropical species are easier to cut than the temperate. Without any 
regard to taxonomic sequence, we shall consider the vegetative 
structure, the strobili, and the prothallia. 

Vegetative structure .—Formalin alcohol, with or without the 
addition of acetic acid, is an excellent fixing agent and, quite contrary 
to prevalent notions, the staining capacity of material seems to improve 
with several months’ immersion. We prefer the following formula: 
formalin 10 c.c., acetic acid 5 c.c., 50 per cent alcohol 100 c.c. 


244 


PTERIDOPHYTES—LY COPODI ALES 


245 


The growing points of stems and roots cut easily in paraffin, and 
when the material becomes too hard to cut in paraffin it can be cut 
without any imbedding. It is easier to get good sections of L. lucidu- 
lum and L. inundatum than of drier species, like L. obscurum and 
L. clavatum. Safranin and Delafield’s haematoxylin is a reliable 
stain. Safranin with anilin blue or light green is also good, and the 
light green gives particularly clear views of the phloem. 

If young sporelings are available they afford a beautiful example 
of a very primitive type of stele, in transverse section showing an 
exarch protostele with 4 or 5 radiating arms of metaxylem, each 
tipped with a comparatively large group of protoxylem cells. In 
most species, this simple radial stele of the sporeling passes into a 
complicated, banded stele in the adult plant. Even in the adult 
plant the protoxylem and metaxylem are easily distinguished in 
sections near the growing point of the stem or root (Fig. 78). Not 
only in Lycopodium, but in any vascular plant, sections at this age 
are useful in pointing out the protoxylem. 

Sections of the stem and root of Lycopodium complanatum, 
mounted on the same slide, show an interesting parallelism of 
structures. Transverse sections of the stem of Lycopodium pithyoides, 
a Mexican species, show not only the stem structures but excellent 
transverse sections of roots which grow down through the cortex. 

The strobilus. —For longitudinal sections, cut a slab from each 
side of the strobilus to insure fixing and infiltration. If a strobilus, 
or similar organ, is simply halved, both pieces are likely to curve. 
Among north temperature species, Lycopodium inundatum is the most 
easily cut. A young strobilus 1 cm. in length may show all stages 
from the archesporium to the spore mother-cell. Iron-haematoxylin 
is the best stain for differentiating the archesporial cells. The divi¬ 
sions in the spore mother-cell stain intensely, so that care must be 
taken not to overstain. 

Strobili of Lycopodium dendroideum or L. obscurum 6 or 7 mm. 
in length show a beautiful series in the development of the sporangium 
from the earliest stages up to tetrads. These young stages fix well 
in chromo-acetic acid, with or without a little osmic acid. 

The gametophyte. —In most species the gametophyte, or prothal¬ 
lium, is subterranean, tuberous, and has no chlorophyll; in other 
species the prothallium is partly subterranean and partly aerial, the 
aerial portion being green and bearing the archegonia and antheridia. 



246 


METHODS IN PLANT HISTOLOGY 


In the third edition of this book, published in 1915, the statement 
was made that no one had yet discovered prothallia of Lycopodium 
in the United States. Two years later, a teacher in the high school 
at Marquette, Michigan, Dr. E. A. Spessard, announced the discovery 



Fig. 78 .—Lycopodium lucidulum: photomicrograph of transverse section of stem near the 
apex. The five deeply staining groups of thick-walled cells are the protoxylem and alternating 
with them are groups of phloem cells; the large, thin-walled cells are metaxylem.- Eastman Com¬ 
mercial Ortho film, Written E filter (orange); arc light; Spencer 4-mm. objective. N.A. 0.65; 
ocular X6; exposure, seconds. Slide by Dr. J. J. Turner, negative bv Dr. P. J Sedgwick 
X360. 

of the prothallia of four species (.Botanical Gazette , 63:51-65, 1917) 
and gave such clear directions for finding prothallia that already 
two papers have appeared, announcing the discovery of prothallia. 
One of these papers, by Dr. Alma Stokey and Dr. Anna Starr 
describes seven stations for prothallia in Massachusetts; and the 


PTERIDOPH YTES—L Y COPODI ALES 


247 


other, by Mr. 0. Degener, adds four more stations in Massachusetts 
and mentions the finding of prothallia near the crater of Kilauea, 
Hawaii. Both papers appeared in the Botanical Gazette of March, 
1924. With these papers describing the finding of prothallia, it is 
probable that prothallia of Lycopodium, especially the older prothallia 
with sporelings attached, will become familiar objects in botanical 
laboratories. 

Once found, the technic is easy. Fix in the special chromo-acetic- 
osmic-acid solution, or this with the acetic reduced to 2 or even to 
1 c.c. Stain in iron-alum haematoxylin and orange for the develop¬ 
ment of archegonia and antheridia. Use safranin, gentian-violet, 
orange for the development of the embryo. For the endophytic 
fungus which stains well with the last-named combination, try the 
method of differentiation of pathogen and host, described at the close 
of the chapter on fungi. 

The prothallia of Lycopodium inundatum , the only species in the 
United States known to have aerial green prothallia, have not yet 
been discovered in this country, although they have been found in 
Europe. 

It would seem natural to get the prothallia by germinating the 
spores, but here again no one has had any notable success, except 
Bruchmann. In some species, the spores do not germinate for several 
years, but when the prothallia are once developed they continue to 
bear archegonia and antheridia for several years. The spores of 
L. Selago germinate in 3 to 5 years after shedding; those of L. clavatum 
and L. annotinum in 6 to 7 years. In L. clavatum and L. annotinum 
archegonia and antheridia develop in 12 to 15 years after the spores 
are shed. L. inundatum germinates more promptly—in 10 days to 
6 months—but no one has succeeded in keeping a culture up to the 
archegonium stage. 

Botanists in Lycopodium localities should look for prothallia. 
Since the subterranean forms are perennial and as large as a grain of 
wheat, some reaching a length of 1.5 cm., it would seem as if they 
should be found wherever Lycopodium grows. 

Selaginella.—-Material of Selaginella, in all phases of the life- 
history, is easy to secure, but not so easy to handle after it is obtained. 
As many as 340 species, mostly tropical, have been described, only 
3 of which are common in the range of Gray’s Manual. Of these 3, 
Selaginella apus is best for sections. It is found in moist or wet 


248 


METHODS IN PLANT HISTOLOGY 


situations on the borders of ponds, along ditches, or on moist meadows. 
While the plant is very small, it has large spores. Several of the 
tropical species are common in greenhouses, and they fruit abundantly. 

Vegetative structure. —Growing points and root-tips are easily 
cut in paraffin. In most species, the older parts of the stem are too 
hard and brittle to cut in paraffin and are too small to cut well free¬ 
hand. It might be worth while to try hydrofluoric acid or the cellulose 
acetate method when that method becomes developed. At present, 
patience and a sharp knife seem to be the only reliance. Some of 
the tropical species, like Selaginella Wildenovii, have stems nearly as 
large as a lead pencil, with polystelic structure, and are not hard to 
cut. The vascular cylinder is an exarch protostele or, when poly¬ 
stelic, each bundle is an exarch protostele. It is exceptionally easy 
to get a brilliant, differentiated stain when once the sections are cut. 

The strohilus. —Very young strobili cut easily in paraffin, but 
after the megaspore coats begin to harden, there are few objects 
which make more trouble than the strobili of Selaginella. For stages 
up to the young megaspores, fix in chromo-acetic acid, with or without 
the addition of a little osmic acid; but for later stages use hot corrosive 
sublimate acetic acid in 50 per cent alcohol. If this is not available, 
use formalin, acetic acohol. Sometimes sections of the difficult 
later stages stay on the slide with Mayer’s albumen fixative. Even 
stages like that shown in Figure 79 should always stay on with Land’s 
fixative. 

The strobili of most species are square in transverse section. 
To get longitudinal sections showing the relations of sporangia, sporo- 
phylls and axis, cut diagonally, from corner to corner, never parallel 
to the flat side. For archesporial cells, use iron-haematoxylin; for 
young megaspores and the development of spore coats, use safranin, 
gentian-violet, orange; for later stages, use safranin and light green. 

The gametophytes. —In most cases the spores germinate while 
still within the sporangium and, in some cases, like Selaginella apus , 
the female gametophyte develops up to the archegonium initial stage 
before shedding. If strobili of this species at the stage shown in 
Figure 79 be broken off and laid down on moist ground so as to keep 
the sporangium moist, dehiscence may not be vigorous enough 
to discharge the megaspores; but development of archegonia and 
antheridia will continue, fertilization will take place, and an embryo 
will be developed while the megaspore is still inclosed within the 


PTERIDOPHYTES—LY COPODI ALES 


249 


sporangium. Such a structure satisfies the definition of a seed. 
Ordinarily, to get the later stages, shake the spore out from the older 
strobili into a Petri dish with the bottom covered by several thicknesses 
of wet filter paper. There is enough nutritive material in the, spores 
to carry them up to the sperm and egg stage. The female gameto- 
phytes within the old spore coats generally orient themselves in the 



Fig. 79 .—Selaginella apus: longitudinal section of strobilus showing a microsporangium 
with germinating microspores on the left; on the right, three of the four megaspores with gameto- 
phytes near the archegonium initial stage; fixed in formalin alcohol, cut in paraffin, and stained 
in safranin and light green; from a preparation by Dr. W. J. G. Land. X80. 


paraffin, the base of the spore being down and the archegonium end 
of the gametophyte being up. A little of some nutrient solution, 
added to the water, will carry the development up to the cotyledon 
stage. The young embryo, with its two cotyledons, its root, and the 
megaspore still attached makes an instructive preparation when 
mounted whole in Venetian turpentine. 

















250 


METHODS IN PLANT HISTOLOGY 


The various stages of the female gametophyte and embryo are 
not hard to stain; but the walls throughout the development of the 
male gametophyte are very thin and extremely hard to stain. 
Safranin and light green is a good combination. Light green in clove 
oil may prove more satisfactory than the alcoholic solution. 

Isoetes. —This peculiar genus is widely distributed and 16 of 
its 64 species occur within the United States. It looks so much like 
a sedge that it is easily overlooked, even when rather abundant. 
As a genus, it is hydrophytic, growing in wet places or even under 
Water. A recent monograph by Dr. Norma Pfeiffer not only gives 
an ecological, morphological, and taxonomic account (with keys in 
English), but gives hundreds of stations, a feature which will enable 
many to find material. 

Vegetative structure— The short, thick stem, even in old plants, 
cuts easily in paraffin. Fix in formalin alcohol and stain in safranin 
and light green. Sporelings with stems about 2 mm. in diameter and 
young plants with stems up to 5 mm. in diameter are best for a 
study of the peculiar vascular system of this plant. These young 
stages fix well in chromo-acetic acid and are not hard to cut. 

Sporangia. —All the sporangia of the plant may be said to con¬ 
stitute a single strobilus of the Selago type. Both longitudinal and 
transverse sections should be cut. The stem is so short that, in 
a plant of medium size, a longitudinal section may include the stem, 
the sporangium, and the sporophyll, up to the top of the ligule. 
Such sections, 10 to 15 n, or even 20 m in thickness, are best for 
demonstration. Transverse sections through the whole cluster of 
sporophylls show the arrangement of megasporophylls and micro- 
sporophylls and also the relations of the sporangia to sporophylls. 

The gametophytes. —The spores are shed in the uninucleate stage, 
and consequently it is not so easy to find the germination as in the 
case of Selaginella. When the large megasporangium begins to 
decay, let the megaspores dry naturally. They retain their power 
of germination for a year at least. Simply wet them with tap water 
and the earlier stages are easily secured, quite clean and ready for 
cutting. There must be soil in the dish for later stages. Try a 
similar method for microspores. Also, look at the top of the stem 
of old plants for stages developing naturally. The cell walls of the 
male gametophyte, as in the case of Selaginella, are rather hard to 
differentiate. Use anilin blue or light green. 


CHAPTER XXII 

PTERIDOPHYTES 

EQUISETALES 

This order was large and prominent in the Carboniferous age, 
but now only a single family, the Equisetaceae, survives. Its only 
genus, Equisetum, contains 24 pieces, 10 of which occur within the 
Gray’s Manual range. Equisetum is often called the “ scouring 
rush,” because the rough stems have been used for scouring kettles. 
The roughness is due to silica. Species, like E. hiemale, which con¬ 
tain much silica, must be treated with hydrofluoric acid before the 
older parts can be cut in paraffin. 

Vegetative Structure. —The roots are very small, but have large 
cells and easily yield good preparations. If a handful of Equisetum 
fluviatile or E. hiemale growing in water be pulled up, scores of root- 
tips may be secured in a few minutes. Fix in chromo-acetic acid 
with a little osmic acid. In case of such small objects it is a good 
plan to add a few drops of eosin to the alcohol during the process of 
dehydrating, in order that the material may be seen more easily. 
The slight staining does no damage, even if more critical stains are 
to be used after the sections are cut. Longitudinal sections of the 
roots may also be obtained by cutting transverse sections of the nodes. 

The growing points of stems may be cut with ease in paraffin. 
E. arvense is particularly favorable on account of the numerous apical 
cells which may be found in a single preparation. 

The “ fertile” stem of Equisetum arvense is so free from silica 
that it can be cut in paraffin without any difficulty. The adult 
vegetative stem of E . arvense , and all stems which contain so much 
silica, must be treated with hydrofluoric acid before imbedding in 
paraffin. However, nearly all of these stems can be cut freehand, 
before fixing, without removing the silica. Fix freehand sections 
in 95 per cent alcohol. Material for paraffin sections should be 
fixed in formalin, alcohol, acetic acid; or in hot alcohol, corrosive 
sublimate acetic. Safranin and anilin blue, with or without a little 
orange, is a good combination. 


251 


252 


METHODS IN PLANT HISTOLOGY 


The Strobilus. — E. arvense affords the most favorable material 
for a study of the development of sporangia, since the strobilus con¬ 
tains almost no silica and, even in its latest stages, is easily cut in 
paraffin. In this species, the young strobili, in the Chicago region, 
can be distinguished from vegetative buds in July; sporogenous 
tissue is well advanced by the middle of August; and the reduction 
divisions occur late in August or early in September (Fig. 80). The 



Fig. 80. Equisetum arvense: A, section of a sporangiophore showing beginning of sporogenous 
tissue, early August condition; B, topography of the strobilus at this stage. A, X580; B, X8. 


spores are not shed until the following April. If you know a patch 
of this species which “fruits” every year, dig up the horizontal 
underground stem in July. The tip of the main axis is almost sure 
to be a strobilus. Dissect away the scale leaves and fix the strobilus 
in chromo-acetic acid with a little osmic acid. August and September 
stages are easy to recognize. If strobili are brought into the labora¬ 
tory in December or January, they shed their spores within a week. 

The spores of Equisetum are excellent for illustrating hygroscopic 
movements. Shake out the spores from the strobili and let them 
dry thoroughly. They can be kept dry for years. When wanted 







PTERIDOPHYTES—EQUISETALES 


253 


for demonstration, put a few on a slide, moisten a little, and watch 
the movements under the microscope. Strobili of other species, like 
E . fluviatile and E. hiemale, contain a large amount of silica and, 
consequently, only the younger stages cut well in paraffin. Hydro¬ 
fluoric acid damages the cell contents more or less. In species like 
these, all stages in the development are found in a single season. 

The Gametophytes. —The spores of Equisetum germinate as soon 
as they are shed, but, like all spores with a considerable amount of 
chlorophyll, they do not long retain the power of germination. A 
comparatively small percentage will germinate a week after shedding, 
and after a month, there may be no germination at all. There is 
no difficulty in growing prothallia to maturity and securing stages 
in the embryo, if fungi or blue-green algae do not appear and ruin 
the cultures. Use Costello’s method for fern prothallia, as described 
on page 263. Consult also a paper by Dr. Elda Walker. 1 This 
paper corrects some previous misconceptions in regard to gameto¬ 
phytes of Equisetum , gives a full account of the gametophyte of 
E. laevigatum, and also directions for finding gametophytes as they 
occur in nature. 

The prothallia fix well in chromo-acetic acid. The younger stages 
may be stained in iron-alum, haematoxylin and mounted in Venetian 
turpentine. The older stages, even of E. arvense , are too large for 
such mounts. E. laevigatum has prothallia a centimeter in diameter. 

For the development of antheridia, the blepharoplast, and the 
development of the sperm, fix in Flemming’s weaker solution and 
stain in iron-haematoxylin. The sperm of Equisetum is the largest 
in Pteridophytes. 

1 Botanical Gazette, 71:378-391, 1921. 


CHAPTER XXIII 

PTERIDOPHYTES 

FILICALES 

This order includes the common ferns and also the Ophioglossaceae, 
which, in the previous edition of this book, was treated as an order, 
Ophioglossales, co-ordinate with Filicales. Some of the ferns are 
sure to be available in almost any locality, and all stages in the life- 
history are easily secured, except early stages in the Ophioglossaceae. 

Vegetative Structure.—From a technical standpoint, the vegeta¬ 
tive structures of Filicales present a wide range of conditions, some 
being so soft that the greatest care must be taken to get them into 
paraffin, while others are so hard that it is almost impossible to cut 
them at all. 

The stem .—Growing points, even of the largest ferns, can be cut 
in paraffin. If the growing point is covered with dense hairs or 
ramentum, either remove the covering entirely or, in case of rather 
fleshy ramentum, remove only the scales which are beginning to turn 
brownish. The white scales will fix and cut. Use chromo-acetic 
acid (1 g. chromic acid and 1 c.c. acetic acid to 100 c.c. water). 
Unless mitotic figures are particularly desirable, it is just as well not 
to add any osmic acid. For illustrating the development of the stem 
from the apical cell, sections 10, 15, or even 20 /* are not too thick. 

Older portions of the stem, or rhizome, in most ferns are easily 
cut while fresh, the sections being transferred to 95 per cent alcohol 
after cutting. But even fairly well-developed rhizomes, after the 
xylem has become lignified sufficiently to stain sharply in safranin, 
can be cut in paraffin, and much finer sections can be obtained than by 
cutting without imbedding (Fig. 81). In digging up rhizomes, do not 
merely dig down until the rhizome can be grasped and then pull it 
up, for such material is sure to show the pericycle of the bundles 
torn away from the parenchyma. Dig carefully around the rhizome 
and then with a very sharp knife cut off pieces which are perfectly 
free. The pieces can be wrapped in wet paper and taken to the labo¬ 
ratory. Then, if they are to be cut without imbedding, cut into 

254 



PTERIDOPHYTES—FILICALES 255 

pieces about 3 cm. long; but material which is to be imbedded should 
be in pieces not longer than 1 cm. 

Pteris aquilina is a good form to practice with. For freehand 
sections, cut as thin as possible, allow 15 to 30 minutes in 95 per cent 


Fig. 81 .—Dicksonia punctilobula: photomicrograph of transverse section of rhizome. Fixed 
in formalin-acetic alcohol, and stained in safranin and light green. Paraffin section 10 n thick. 
Eastman Commercial Ortho film, Wratten E filter (orange); arc light; J. Swift and Son 1-inch 
lens; exposure, 1 second. Negative by Dr. P. J. Sedgwick. X33. 

alcohol, transfer to 50 per cent alcohol and stain over night in safranin. 
Rinse in 50 per cent alcohol and stain in light green. Paraffin sections 
2 or 3 cm. back of the apex are very instructive, for only the proto- 
xylem will be sufficiently lignified to stain red with safranin, the 





256 


METHODS IN PLANT HISTOLOGY 


metaxylem still being thin walled and staining with the light green. 
This rhizome affords an excellent illustration of a mesarch polystele. 

Dicksonia punctilobula has a small rhizome, often on the surface 
of the soil or rock, so that it is easy to get good clean pieces. About 
2 or 3 cm. back of the growing point, the xylem is well lignified, but 
the material still cuts well in paraffin. One could hardly find a 
better illustration of a mesarch amphiphloic siphonostele (Fig. 81). 



Fig. 82. —Botrychium obliquum: photomicrograph of transverse section of rhizome. Fixed 
in formalin-acetic acid-alcohol, and stained in safranin, gentian-violet, orange. Paraffin section, 
10 n. Eastman Commercial Ortho film, Wratten E filter (orange); arc light; J. Swift and Son 
1-inch lens; exposure, 1 second. Negative by Dr. P. J. Sedgwick. X45. 

Botrychium is widely distributed but individual plants are not 
abundant. The stem is erect, subterranean, and has an endarch 
siphonostele with secondary wood. Trim away the roots, which are 
very thick and fleshy in B. obliquum , fix in formalin-alcohol-acetic 
acid, and imbed in paraffin. Even the older parts of old stems can 
be cut in paraffin if you are sufficiently careful (Fig. 82.) Transverse 
sections from the base of the bud down to the secondary wood will 
give a beautiful series in the development of the stele. 

The bud at the top of this rhizome is an interesting object. The 
leaf is in its fourth year when it appears above ground, and, conse- 









PTERIDOPH YTES—FI LIC ALES 


257 



quently, the bud contains young leaves of three successive seasons. 
Two of the three show a differentiation into sterile and fertile portions. 

In Osmunda, and in many other ferns of similar habit, the 
rhizome is surrounded by the very hard leaf bases. Good sections 
of the central cylinder can be secured only by dissecting away these 
hard leaf bases and any hard portions of the cortex before attempting 
to cut sections. A short dis¬ 
tance back of the growing 
point will be found a region 
which will show practically all 
the structures of the mature 
stem, which will be easy to 
cut. Even in this region the 
leaf bases should be dissected 
away. From the apical cell 
back to the region where the 
sclerenchyma is beginning to 
turn brown, the material is 
easily cut in paraffin. Older 
portions should be cut free¬ 
hand. Osmunda affords an 
excellent illustration of the 
mesarch siphonostele (Fig. 83). 

The rhizome of Adiantum 
affords a good illustration of 
leaf gap and leaf trace. The 
vascular cylinder is a mesarch 
siphonostele; but there are few 
sections like Figure 81, because 
the clyinder is so interrupted 
by leaf gaps. This rhizome 
cuts well without imbedding. The petiole of Botrychium , in trans¬ 
verse sections below the fertile spike, shows the interesting leaf-trace 
situation which proves that the fertile spike consists of a pair of 
pinnae fused together. 

The ferns of the Gray’s Manual range afford no very satisfactory 
material for illustrating the protostele, although protosteles occur 
in Lygodium and Trichomanes. The most satisfactory material is 
Gleichenia, a very common and very beautiful fern in tropical and 


Fig. 83 .—Osmunda cinnamomea: photomicro¬ 
graph of single bundle of the mesarch siphonostele. 
Stained in safranin and anilin blue. Eastman Com¬ 
mercial Ortho film, Wratten C (blue-violet) filter; 
Bausch and Lomb 4-mm. objective, N.A. .65; ex¬ 
posure, 4 seconds. Negative by Dr. P. J. Sedg¬ 
wick. X108. 




258 


METHODS IN PLANT HISTOLOGY 



subtropical regions, but almost never seen in greenhouses nor even in 
botanical gardens. Formalin alcohol material is easily cut without 
imbedding and is easy to stain. 

The stems of tree ferns require special treatment. With the 
large leaf bases partly cut away with a sharp razor, transverse sec¬ 
tions are easily cut for a con¬ 
siderable distance below the 
apex. Material fixed in for¬ 
malin alcohol cuts very well. 
If fresh material is to be cut, 
the softer portions should be 
flooded with alcohol after each 


section. Farther down, there 
will be a region where sections 
can be cut without any flood¬ 
ing, and still farther down, it 
will be difficult or impossible 
to cut sections across the whole 
stem. Sections 1 or 2 cm. 
thick, cut smooth on the ends, 
may be kept in 95 per cent 
alcohol or in glycerin in large 
glass dishes of the Petri dish 
pattern. Better still, clear 
such sections in xylol and pre¬ 
serve in cedar oil. 

The root .—The roots of 
Filicales develop from a strong 
apical cell. For mitotic fig¬ 
ures and the development of 
the root from the apical cell, 
fix the tip in chromo-acetic acid with a little osmic acid. If the 
development of the root is the principal object, stain in safranin and 
light green, or in the safranin, gentian-violet, orange combination; 
if mitotic figures are to be studied, stain in iron-haematoxylin with 
a very light counter-stain in orange. The comparatively large root- 
tips of Botrychium are excellent for the apical cell and its segments. 
Dicksonia punctilohula can also be recommended; but even the very 
small root-tips of most of our ferns will yield good preparations. 


Fig. 84. —Angiopteris evecta: photomicrograph 
of a transverse section of a root, showing the 
polyarch, exarch siphonostele. Eastman Commer¬ 
cial Ortho film, Wratten E (orange) filter; arc light; 
J. Swift and Son 1-inch lens; exposure, f second. 
Negative by Dr. P. J. Sedgwick. X25. 





PTERIDOPHYTES—FILICALES 


259 



Roots of tree ferns are sometimes available in greenhouses. In 
some species the stem is covered by a dense felt of small roots, some 
of which will be white and soft at the tip. These roots are likely to 
have about the diameter of onion root-tips, and the beauty of prepara¬ 
tions made from them could hardly be excelled. In the tropics, where 
the plants are often in the 
spray of cataracts and the 
lower part of the trunk is often 
washed by mountain streams, 
a thousand tips might be se¬ 
cured from a single specimen. 

The older roots of Botrych- 
ium, especially the large fleshy 
roots of B. obliquum, cut very 
easily and show a simple exarch 
protostele with 4 or 5 proto- 
xylem points. 

The roots of Angiopteris 
and Marattia, which become as 
large as a lead pencil, may be 
secured in some greenhouses. 

They cut easily after fixing in 
formalin alcohol and furnish a 
fine example of the exarch pro¬ 
tostele, common to all roots 
(Figs. 84 and 85). 

The structure of the leaf will 
appear in sections cut to show 
the sporangia. 

The Sporangia. —To illus¬ 
trate the character of the an¬ 
nulus, select sporangia which are just beginning to turn brown. Fix 
in formalin alcohol and dehydrate as if for paraffin sections; after the 
absolute alcohol, transfer to 10 per cent Venetian turpentine. Stain¬ 
ing is neither necessary nor desirable. 

The various relations of sorus and indusium are best illustrated 
by rather thick sections (10 to 20 y) of material in which the oldest 
sporangia have barely reached the spore stage. Fix in formalin 
alcohol and stain in safranin and anilin blue. 


Fig. 85.— Angiopteris evecta: photomicrograph 
showing a detail of the stele shown in Figure 84. 
Stained in safranin. Eastman Commercial Ortho 
film, Wratten E (orange) filter; Bausch and Lomb 
8-mm. objective, N.A. .50; exposure, lj seconds. 
Negative by Dr. P. J. Sedgwick. X60. 









260 


METHODS IN PLANT HISTOLOGY 


For the development of sporangia use hot corrosive sublimate- 
acetic acid-alcohol, or chromo-acetic acid (1 g. chromic acid and 2c.c. 
acetic acid to 100 c.c. of water). A larger proportion of figures will 
be secured by adding 5 or 6 drops of 1 per cent osmic acid to this 
solution. Sections should not be more than 10 u in thickness and, 
for mitotic figures, 5 y is thick enough. 



Fig. 86. Osmunda cinnamomea: photomicrograph of sporangia with spore mother-cells 
in various stages of division; fixed in Flemming’s weaker solution and stained in iron-alum haema- 
toxylin; from a preparation by Dr. S. Yamanouchi. Negative by Miss Ethel Thomas. X114. 

For the reduction of chromosomes, the sections should not be 
thicker than 5 y. Osmunda is particularly good for this purpose be¬ 
cause the number of chromosomes is comparatively small. The 
young sporangia of Osmunda cinnamomea and 0 . Claytoniana show 
the mother-cell stage in the autumn, but the division into spores does 
not occur until the following spring, in the vicinity of Chicago, the 
mitotic figures being found during the latter part of April (Fig/86). 
O. regalis does not reach the mother-cell stage in the autumn. Mate¬ 
rial for mitosis should be collected during the first two weeks in May. 




PTERIDOPHYTES—FILICALES 


261 


Various species of Pteris are common in greenhouses and are very- 
good for development of sporangia. Any fern of the Aspidium 
type will yield a good series, and some, like Cyrtomium, may show a 
fine series in a single sorus. Marattia, which is likely to be found in 
botanical gardens, will illustrate the “synangium” type; Angiopteris 
has a sporangium which forms an easy transition to that of the 
Cycadales. 

For the development of a typical sporangium of the eusporangiate 
type, nothing is better than Botrychium. Buds of B. virginianum 
taken in September or October show sporangia with well-marked 
sporogenous tissue. For a study of the development of sporangia, 
cut off the fertile portion and fix it separately, using chromo-acetic 
acid with 5 or 6 drops of 1 per cent osmic acid to 100 c.c. of the solu¬ 
tion. Stain in iron-alum haematoxylin. The reduction divisions in 
the spore mother-cell take place after the leaf arrives above the surface. 

The Prothallia.— Prothallia can usually be found on the pots in 
the ferneries of greenhouses. Ripe spores of some fern or other can 
be obtained at any greenhouse at any time in the year, and spores 
of most of our native ferns germinate well and produce good prothallia, 
even if the sowing is not made for several months after the spores 
have been gathered. 

Fine prothallia of Pteris aquilina have been grown two years 
after the spores were gathered. Some, however, must be sown at 
once, or they will not germinate at all. Spores which are large and 
contain enough chlorophyll to make them appear greenish should 
be sown at once. The spores of the common Osmunda regalis , and 
of the other members of the genus, must be sown as soon as ripe, or 
they fail to germinate. The prothallia of 0. regalis } if carefully 
covered with glass, may be kept for a long time, and they become 
quite large. Prothallia of this fern in the writer’s laboratory pro¬ 
duced ribbon-like outgrowths 5 mm. wide and more than 5 cm. in 
length. These prothallia continued to produce archegonia, anther- 
idia, and ribbon-like outgrowths for more than a year, when they 
suddenly “ damped off.” Lang watered prothallia with a weak 
solution of permanganate of potash, which kills the fungi but does 
not injure the prothallia. He does not state the strength of the 
solution, but 4 or 5 crystals to 1 liter of water seems to be effective. 

The prothallia of most ferns will grow for a long time under such 
conditions. Pteris aquilina and many other ferns often furnish a 


262 


METHODS IN PLANT HISTOLOGY 


good supply of antheridia 3 weeks after sowing, and the archegonia 
appear soon after, but it is well to make sowings 6 weeks before 
material is needed for use. In P. aquilina and in many others, if 
the spores are sown too thickly only antheridial plants will be ob- 



Fig. 87 .—Pteris aquilina: A, filamentous stage; B, the apical cell has been established and 
several segments have been cut off; the figure shows the initial rhizoid and also three rhizoids 
coming from the main body of the prothallium; C an older prothallium covered with antheridia in 
various stages of development; from a drawing by Miss M. E. Tarrant. 

tained (Fig. 87). If prothallia are to produce archegonia, they must 
have sufficient room and nutrition. 

The best method we have ever seen for growing fern prothallia 
was devised by Mr. M. Costello, head gardener at the University of 
Chicago. The diagrammatic Figure 88 will make the method clear. 
Select a clean flower-pot, as porous as possible, and pack it full of 
wet Sphagnum. Wet the outside of the pot and invert it in a pan of 







PTERIDOPHYTES—FILICALES 


263 


water. Sow the spores on the surface of the pot and cover with a 
bell jar. No further wetting is necessary, except to take care that 
the water in the pan does not dry up. With Pteris longifolia, there 
may be antheridia in 2 weeks; archegonia in 3 or 4 weeks; and in 
5 or 6 weeks, abundant sporophytes in various stages. Prothallia 
grown by Costello’s method are entirely free from soil and, conse¬ 
quently, very convenient for cutting or for mounting a hole. 

While there should always be a study from living material, it is 
worth while to make permanent mounts, even for habit study. For 
such study, the prothallia 
should be mounted whole. 

Fix in the special chromo- 
acetic-osmic-acid solution. 

If the material shows any 
tendency to break up, use 

2 c.c. of acetic acid instead of 

3 c.c. In cities where water is 
treated with copper or other 
substances, the difficulty may 
sometimes be due to the water 
rather than to any excess of 
acetic acid. Formalin-acetic 
acid (10 c.c. formalin, 5 c.c. 
acetic acid to 100 c.c. water) 
is good for material which is 
to be mounted whole. Stain 
some in iron-alum haematoxy- 
lin, and some in Magdala red 
and anilin blue. Mount in 
Venetian turpentine, using 
material from each stain for each mount. Select stages so that 
each preparation will show the filamentous stage, the apical cell stage, 
the group of initials stage, and also antheridia and archegonia. 

For sections, the chromic series is better than formalin or cor¬ 
rosive sublimate. If the gradual processes of dehydrating, clearing, 
and infiltrating have been carefully observed, about 15 or 20 minutes 
in the bath should be sufficient. About 10 /* is a good thickness for 
such views as are shown in Figures 89, 90, 91. Safranin, gentian- 
violet, orange is a good stain. 



prothallia. 










264 


METHODS IN PLANT HISTOLOGY 


For the development of the antheridium and sperm, and especially 
for the blepharoplasts and their transformation, cut about 3 y in 
thickness and use iron-haematoxylin; for the development of arche- 
gonia, cut at 5 y and stain in the safranin, gentian-violet, orange 
combination. 

The gametophyte of Botrychium is subterranean and tuberous. 
It sometimes reaches a length of 7 to 12 mm. and a thickness of 4 to 
5 mm. Usually, it is not more than 5 or 6 mm. long and 2 or 3 mm. 
thick. Gametophytes showing the development of antheridia and 



Fig. 89. —Osmunda cinnamomea: photomicrograph of vertical section from the notch toward 
the base of the prothallium showing four stages in the development of the archegonium—chromo- 
acetic acid; safranin, gentian-violet; from a preparation by Dr. W. J. G. Land. Negative by 
Miss Ethel Thomas. X425. 

archegonia are not likely to be more than 2 or 3 mm. long and 1 or 
2 mm. thick. Near large plants, look for small sporelings, not more 
than 1 or 2 cm. in height. Dig very carefully and you may find the 
gametophytes attached. The soil should be examined for smaller 
specimens. Most of the gametophytes will be found at a depth of 
1 to 3 cm. Fix in chromo-acetic acid. 

No one has yet succeeded in raising the prothallia from the spores. 
The prothallia always contain an endophytic fungus, but even when 
this is present no one has succeeded in raising prothallia. 

The prothallium of Ophioglossum is harder to find or, perhaps 
it would be better to say, harder to recognize, for it is also subter- 




PTERIDOPHYTES—FILICALES 


265 


ranean and tuberous and, besides, looks so much like the root that 
you may not recognize it, even when you have it in your hand. 

The Embryo. —Instructive mounts of the whole embryo, with the 
pro thallium still attached can be made by the Venetian turpentine 
method. Iron-alum haematoxylin, with the stain not too deep, 



Fig. 90. — Osmunda cinnamomea: 
photomicrograph of vertical section of 
prothallium with an early stage in the de¬ 
velopment of the archegonium, showing the 
basal cell, two neck cells, and, between 
them, the cell which is to give rise to the 
neck canal cell, the ventral canal cell, and 
the egg—chromo-acetic acid; safranin, 
gentian-violet; from a preparation by Dr. 
W. J. G. Land. Negative by Miss Ethel 
Thomas. X425. 



Fig. 91. —Osmunda cinnamomea: 
photomicrograph of a vertical section with 
a young archegonium, showing the neck 
canal cell with two nuclei, the ventral 
canal cell, the egg, and the basal cell— 
chromo-acetic acid; safranin, gentian- 
violet; from a preparation by Dr. W. J. G. 
Land. Negative by Miss Ethel Thomas 
X293. 


is good; Magdala red and anilin blue are more transparent and will 
show the structure of the root, if the two stains are well balanced. 

For sections, cut longitudinally, perpendicular to the prothallium. 
Pteris longifolia may show the young embryo within 3 or 4 weeks; 
Osmunda , somewhat later. 






266 


METHODS IN PLANT HISTOLOGY 


Nepkr odium is generally parthenogenetic. The embryos, devel¬ 
oped while the prothallia are still very small, come from tissue cells of 
the prothallium. Pteris cretica is frequently parthenogenetic. 

The Heterosporous Filicales. —'The four genera, Pilularia , Mar- 
silia, Salvinia, and Azolla are aquatic, the first two growing rooted 
but more or less submerged, and the other two floating freely on 
the water. Marsilia is the most available and convenient laboratory 
type of this group. It is easily grown in a pond or in an aquarium 
in the greenhouse. In setting it out in a pond, select a place with 
a gently sloping bank, so that part of the material may be under 
water and part may creep up the bank. In the greenhouse, a rec¬ 
tangular aquarium may be tilted to secure the same conditions. 
The portions which are not under water will continue to fruit during 
the summer and autumn. The whole sporocarp cuts easily in paraffin 
during the development of sporangia, the division of the spore 
mother-cells, and even during the earlier stages in the formation of 
spores. Except in the case of the youngest sporocarps, it is better 
to cut off a small portion at the top and at the bottom to facilitate 
fixing and infiltration. The mother-cell stage and the young spores 
will be found in sporocarps which are just beginning to turn brown. 
In nature, no further nuclear divisions take place within the sporan¬ 
gium until the next spring, but the wall of the sporocarp becomes 
extremely hard. Sporocarps for germinating should not be collected 
until they are so hard that it is impossible to crush them between the 
thumb and finger. They can be kept in a box until needed for use. 
When you find them in good condition, make a big collection, for 
they retain their power of germination almost indefinitely, sporocarps 
from poisoned herbarium specimens 50 years old germinating readily. 
Sporocarps which have been kept in 95 per cent alcohol for years ger¬ 
minate almost as quickly as those which have been kept in a dry box. 

To germinate sporocarps, cut away a portion of the hard wall along 
the front edge and place the sporocarp in a dish of water. The 
gelatinous ring with its sori will sometimes come out in a few minutes. 
In less than 24 hours, sometimes within 10 or 12 hours, microspores, 
starting from the one-cell stage, will produce the mature sperms; 
and the development of the female gametophyte is equally rapid. 
Starting with the uninucleate megaspore, the stage found when the 
gelatinous ring comes out, the archegonium may be developed and 
fertilization may occur within 12 hours; and within 36 hours, stages 


PTERIDOPHYTES—FILICALES 


267 


like that shown in Figure 92 may be reached. At the end of a week, 
there may be green sporophyte more than a centimeter in length. 

It is obvious that material should be fixed at frequent intervals 
if one is to secure a series of stages in the development of the gameto- 
phytes and embryo. 

For sections showing the development of the antheridium and 
sperms, it is better to remove the megaspores from the sorus, since 
they occasion considerable difficulty in cutting. Fix in the special 
chromo-acetic-osmic-acid solution, cut 5 /jl thick, and stain in iron- 
alum haematoxylin. 



Fig. 92 .—Marsilia quadrifolia: upper portion of megaspore with an archegonium containing 
a young embryo. X212. 

The older megaspores, with the archegonium or embryos at the 
apex, are very hard to cut. Fix in hot alcoholic corrosive sublimate- 
acetic, or in alcohol-formalin-acetic acid. It will facilitate infiltration 
and cutting it you prick each megaspore with a sharp needle before 
fixing. Since the archegonium is at the apex of the megaspore, 
the pricking need do no damage. If sections come loose from the 
slide, use Land’s fixative. Cut 5 to 10 ju thick and stain in safranin, 
gentian-violet, orange. 

The sperm, which in Marsilia has an unusually large number 
of turns in the spiral, is easily mounted whole. When the sperms 
have become numerous, put several megaspores upon a slide and 
heat gently until dry. Then wet the preparation in any alcohol and 
stain sharply in acid fuchsin. Dehydrate in absolute alcohol for 



268 


METHODS IN PLANT HISTOLOGY 


at least 10 minutes, clear in clove oil, and mount in balsam. Such 
a preparation will often show a score of sperms in the gelatinous 
funnel leading down to the neck of the archegonium. 

Azolla is not difficult to obtain, and it is easy to get a series of 
stages in the development of the micro- and megasporangia; but 
it is not at all easy to find the gametophytes, since the spores ger¬ 
minate only after they have been set free by the decay of the plant. 
Azolla does not fix well in any of the chromic-acid series, because it 
catches so much air that it will not sink. Alcohol-formalin-acetic, 
or hot alcoholic corrosive sublimate-acetic acid, formalin (4 g. 
corrosive sublimate, 5 c.c. formalin; 5 c.c. acetic acid, 100 c.c. of 
50 per cent alcohol) can be recommended for both Azolla and Salvinia. 
Both of these forms grow well in the greenhouse, floating on the water 
in tanks or aquaria; but Azolla seldom fruits under such conditions. 
Salvinia sometimes produces microsporangia in the greenhouse, but 
megasporangia are comparatively rare. 


CHAPTER XXIV 


SPERMATOPHYTES 

Material in this group ranges from structures so delicate that 
they require more skill and patience than the coenocytic algae to 
structures so hard that the method for rock-sections is the best way 
to get preparations. We cannot hope to give even approximately 
complete directions for making preparations, but must be content 
to give a few hints which may prove helpful in collecting material 
and in securing mounts of the more important structures. We 
shall consider the gymnosperms and the angiosperms separately, 
although in many respects the technic is the same for both. 

GYMNOSPERMS—CYCADALES 

Cycas revoluta, the Sago Palm, can be found in almost any large 
greenhouse which keeps decorative plants. The large conservatories 
of city parks may keep, in addition, some species of Ceratozamia or 
Encephalartos. Only one cycad, Zamia , occurs in the United States, 
and it is confined to Florida. In Encephalartos and Ceratozamia the 
development of the ovule, and even the development of the female 
gametophyte up to the fertilization period, takes place quite naturally 
in the greenhouse, where pollination is not likely to occur; but in other 
genera, the female cones, or at least their ovules, nearly always 
abort unless fertilization takes place. The vegetative structures are 
natural enough, but, with the exception of leaves and small roots, are not 
so available, since material of the stem would mean damage to the plant. 

The Vegetative Structures.—All the vegetative structures cut 
rather easily. 

The stem.—Zamia / which grows in various parts of Florida, is 
the most available material. Directions for handling the stem are 
given on page 132. 

Stems of the larger cycads are not likely to be obtained, except 
in the field, and they are confined to tropical and subtropical regions. 

i Material of Zamia can be obtained at $1.00 a plant (express, collect) by addressing the 
Plant Pathologist, Agricultural Experiment Station, University of Florida, Gainesville, Fla. This 
is not a commercial matter, but an accommodation to botanists. 

269 


270 


METHODS IN PLANT HISTOLOGY 


They cut better while fresh; consequently, if one can get material, 
it is a good plan to send it to the laboratory and have it cut before 
fixing. Even transverse sections are not difficult to cut while 
fresh (Fig. 93). A piece of cycad trunk 15 to 30 cm. in diameter 
and 20 cm. in length will survive a journey of 6 weeks or even 2 
months, if care be taken to coat the exposed ends with a mixture 
of melted paraffin and moth balls, using 3 or 4 moth balls as large 
as marbles to half a kilo of paraffin. If material is to be fixed before 
cutting, use 6 to 10 per cent formalin in water. 


Fig. 93.— Dioon spinulosum: photomicrograph of transverse section of wood 
sum, cut from fresh material. X105. 


of Dioon spinulo- 


Dr. La Dema Mary Langdon succeeded in cutting paraffin sections 
of the adult wood of Dioon spinulosum. 1 She fixed 1 to 2 cm. cubes 
of adult wood in formalin-acetic acid-alcohol (6 c.c. formalin, 
3 c.c. acetic acid, 100 c.c. of 50 per cent alcohol). After thorough 
washing, 24 to 48 hours in running water, the blocks were softened for 
3 to 6 weeks in 50 per cent hydrofluoric acid in water. With dry 
material, the cubes were boiled and cooled repeatedly to remove air. 
The usual gradual processes of dehydrating, clearing, and infiltrating 

1 Botanical Gazette, 70:82-84, 1920. 



'•••Iff 

Mm 























SPERMATOPHYTES—GYMNOSPERMS 


271 


were then followed and sections were cut with a sliding microtome, 
with the knife placed obliquely, as in cutting celloidin. 

The course of the vascular bundles, as they pass to the cones, is 
quite peculiar. Instructive preparations may be made by cutting 
longitudinal sections, about 3 mm. thick, through the apex of the 
stem and, without staining, clearing thoroughly and mounting in 
balsam. In this way we have mounted sections 5 cm. long, 15 cm. 
wide, and 3 mm. thick. 

The course of the bundles in the xylem zone and in the cortex may 
be traced by clearing the cubes in xylol and then transferring to 
equal parts of xylol and carbon disulphide. Placed on a glass plate 
with an electric light bulb beneath, the bundles are quite distinct. 

The root .—Small roots, up to a centimeter in diameter, are easily cut 
freehand. The tender root-tips and also the peculiar “ root-tubercles” 
should be fixed in chromo-acetic acid and imbedded in paraffin. 

The leaves .—The young tender leaves should be fixed in formalin 
alcohol and imbedded in paraffin. The adult leaves are rigid and 
cut well freehand. It is a good plan to tie several leaflets together 
with a string and then cut across, about a centimeter beyond the 
tied portion, so that the whole will be like a leaflet 5 or 6 mm. thick. 
Dip the whole thing in paraffin two or three times. Of course, there 
is no infiltration but the paraffin holds the leaflets in place. Cut in 
a sliding microtome with an oblique stroke. The sections fall out 
from the paraffin, which is easily skimmed away. Fix the sections 
for an hour in 95 per cent alcohol, stain in safranin and light green, 
clear in clove oil, transfer to xylol, and mount in balsam. 

Spermatogenesis. —Except in the earliest stages, the staminate 
cones are too large to be cut whole. The individual sporophylls, 
with their sporangia, cut easily up to the formation of microspores; 
then the sporangium wall hardens rapidly and cutting becomes 
difficult. Up to the young microspore stage, fix in chromo-acetic- 
osmic-acid solution (1 g. chromic acid, 1 c.c. acetic acid, 1 c.c. 1 per cent 
osmic acid to 100 c.c. water). With a larger proportion of acetic 
acid, our results have not been satisfactory. Fix later stages 
in formalin-acetic acid-alcohol. Transverse sections are more 
instructive and are more easily cut, since the peripheral end of the 
sporophyll can be cut only in younger stages. In all the genera of 
cycads, the microspore germinates while still within the sporangium, 
the pollen grain at the time of shedding consisting of a prothallial 


272 


METHODS IN PLANT HISTOLOGY 


cell, a generative cell, and a tube cell. For preparations at the 
shedding stage, shake the cone over a piece of paper and pour the 
pollen into water. After 15 or 20 minutes, put it into the fixing 
agent. In wind-pollinated plants, the pollen would look shriveled 
if you put it into a fixing agent before it regained its turgidity. 

The pollen, mounted whole, makes beautiful preparations. Fix 
in formalin-acetic acid (formalin 10 c.c., acetic acid 5 c.c., water 



Fig. 94.— Ceratozamia mexicana: A, pollen grain which has been in a sugar solution for 
two days; X876; B, nucellus with numerous pollen tubes; X17; C, basal end of pollen tube 
showing the persistent prothallial cell; outside it the stalk cell; and, above, the two sperms still 
inclosed in the sperm mother-cells; X156. 

100 c.c.). Stain in iron-alum haematoxylin and follow the Venetian 
turpentine method. Carmalum, as described in the Power’s methods 
on page 178, is also satisfactory. If some material is put into a 
5 or 10 per cent sugar solution for two or three days, early stages in 
the formation of the pollen tube may be added. In a week or two 
the tubes may reach several times the length of the pollen grain; 
but, as far as we know, the generative cell has never divided in culture 
solutions (Fig. 94A). 








SPERMATOPHYTES—GYMNOSPERMS 


273 


The development of the pollen tube and its structures must be 
studied in sections of the nucellus. As soon as the integument is 
removed the nucellus is exposed and the position of the pollen tubes 
is easily determined, since the haustorial portions of the tubes form 
brownish lines radiating from the nucellar beak. Having learned 
the location of the pollen tubes, it is better not to remove the integu¬ 
ment, but to remove the female gametophyte; then cut from the 
underside of the nucellus against the hard, stony layer of the integu¬ 
ment so as to remove a small piece of the nucellus 5 to 7 mm. square, 
according to the species. Fix in chromo-acetic-osmic acid (1 g. 
chromic acid, 2 c.c. acetic acid, 1 c.c. 1 per cent osmic acid to 100 c.c. 
water). Nothing surpasses iron-alum haematoxylin for all the 
stages in the development of the male gametophyte (Fig. 94 A-C). 
By using a 2 per cent iron-alum for 5 or 6 hours and staining over 
night, or even 24 hours, the stain can be drawn so precisely that the 
portion of the cilia between the blepharoplast and the surface can be 
differentiated from the free portion (Fig. 95). 

The pollen tubes, with their sperms, make instructive prepara¬ 
tions when mounted whole. Fix the nucellus, with its pollen tubes, 
as if for paraffin sections. About 6 per cent formalin in water has 
proved successful. Wash in water for half an hour and stain in 
aqeous safranin, 1 or 2 hours. Extract the stain until it is satis¬ 
factory, and then transfer to 10 per cent glycerin and follow the 
Venetian turpentine method. When the turpentine becomes thick 
enough for mounting, tease the pollen tubes from the nucellus and 
mount with pieces of cover-glass under the cover to prevent crushing. 
Iron-alum haematoxylin, which is so satisfactory for staining sections 
of sperms, is the most unsatisfactory stain we have tried for staining 
the cycad pollen-tube structures whole. 

The immense sperms of the cycads, more than 100 ix in diameter, 
can be seen with the naked eye. If the tip of the nucellus with the 
pollen chamber be removed when the sperms are mature, the large 
pollen tubes, 300 to 500 m in diameter, are very conspicuous. Even 
with the naked eye, the movements of the sperms can be seen; and 
with a pocket lens one can see some details. Place the piece of 
nucellus on a slide in a drop of water and examine with a 16 mm. 
objective. The amoeboid movements of the sperm and also a quick 
movement reminding one of the sudden jerky movement of Vorti- 
cella, are easily seen; and, by closing the diaphragm a little, some 


274 


METHODS IN PLANT HISTOLOGY 


movement of the cilia should be distinguishable. Zamia is the only 
genus available in the United States; but even its rather small female 
cones, as they near the fertilization stage, will keep 4 or 5 weeks 
after they have been removed from the plant. 



Fig. 95 .—Ceratozamia mexicana: photomicrograph of a longitudinal section of a sperm, 
showing the very large nucleus, thin sheath of protoplasm which is thicker at the apical end, and 
blepharoplast cut across in various places and bearing numerous cilia. At this stage, the sperm is 
swimming in the pollen tube. Cramer, contrast plate. Negative by Miss Ethel Thomas. X506. 

In the region of Miami, Florida, pollination takes place late in 
December or early in January; blepharoplasts appear in March, and 
swimming sperms are found during the first 2 weeks of June. In 










SPERMATOPHYTES—GYMNOSPERMS 


275 


northern Florida, all stages are 2 or 3 weeks later. If one is so for¬ 
tunate as to get cones of Ceratozamia, the sperms should be found from 
the last week in June through the second week in July. 

Oogenesis.— The ovules of the Cycads and Ginkgo are very large, 
and, when mature, thin sections cannot be cut by any method yet 
discovered. In younger stages it is not difficult to get good sections 
of the entire ovule. Slabs should be cut from two sides of the ovule 
to facilitate fixing and infiltration. During free nuclear stages in the 
endosperm, and even during earlier stages in the formation of walls, 
care must be taken that the slabs may not cut into the endosperm, 
or even too near to it; for the endosperm is so turgid that it may 
even break out, and, at least, will be distorted. Even after the ovule 
approaches its full size, it can be cut entire, until the stony layer 
begins to harden. Paraffin sections of the entire ovule, cut 15 to 
20 \x thick, and stained rather lightly in safranin, gentian-violet, 
orange, make very instructive preparations. When the fresh ovule 
can no longer be cut easily with a razor, it is not worth while to try 
to cut it in paraffin. Interesting preparations may be made by 
cutting from the median longitudinal portion of the ovule a slab 
about 5 mm. thick. The slab should be fixed, washed, dehydrated, 
and cleared in xylol. It should then be kept in a flat-sided bottle. 
Such a preparation shows the integument, micropyle, nucellus with 
its beak, pollen tubes, the stony and fleshy layers, general course 
of vascular bundles, and the female gametophyte with its archegonia. 

For thin sections of the archegonia, a cubical piece with an edge 
of 6 or 8 mm. should be cut from the top of the endosperm with 
a very sharp, thin blade. The slightest pressure upon the archegonia 
will ruin the preparations. 

Chromo-acetic-osmic acid (1 g. chromic acid, 2 c.c acetic acid, 
and 1 c.c. of 1 per cent osmic acid to 100 c.c. of water) is a good 
fixing agent for all stages in oogenesis. During the free nuclear stage 
and early wall stage in the female gametophyte, some plasmolysis 
is to be anticipated. Hot alcoholic corrosive sublimate-acetic acid 
sometimes fixes these stages with less distortion. 

Sporophyte.—During the period of simultaneous free nuclear 
division, which follows the fertilization of the egg, the mitotic 
figures are quite striking and are easily stained (Fig. 96). 

After the embryos begin to grow down into the endosperm, 
oblong pieces containing the embryos should be cut out. 


276 


METHODS IN PLANT HISTOLOGY 


After the cotyledons appear, useful preparations may be made by 
dissecting out the entire embryos, which may be fixed in chromo- 
acetic acid, washed, stained in eosin or in Delafield’s haematoxylin, 
placed in 10 per cent glycerin, and mounted by the Venetian turpen¬ 
tine method. Since the suspensors become long and irregular, each 
embryo should be placed in a separate dish, lest the suspensors be¬ 
come entangled and broken. 



Fig. 96. —Zamia floridana: photomicrograph of a small portion of a proembryo showing 
simultaneous free nuclear division. Fixed in chromo-acetic-osmic acid and stained in safranin, 
gentian-violet, orange. Cramer contrast plate; Spencer 4-mm. objective, N.A. .65; ocular X4; 
yellowish-green filter; camera bellows, 50 cm.; arc light; exposure, 6 seconds. Negative by Miss 
Ethel Thomas. X413. 

After the stony layer becomes hard, it is better to use a small 
fret saw for opening the ovule. Before the embryo has pushed down 
into the endosperm, the ovule should be sawed in two transversely. 
The endosperm and nucellus can then be picked out and treated as 
desired. After the tip of the embryo reaches the middle of the endo¬ 
sperm, the ovule should be sawed open longitudinally. 

GYM NOSPERMS—GINKGOALES 

From the standpoint of technic, the Ginkgoales, now represented 
only by Ginkgo biloba, are less difficult than the Cycadales, but the 
difficulties are somewhat similar. 

The Vegetative Structures— The adult stem is harder to cut than 
Pinus, but good sections should be secured by boiling in water and 
soaking for a few days in glycerin. Transverse sections of the “ spur ” 
shoots are easily cut. They have a comparatively large pith and 
narrow zone of wood, thus contrasting sharply with a long shoot of 
the same diameter, which has a small pith and wide zone of wood. 











SPERMATOPHYTES—GYMNOSPERMS 


277 


The petiole of the leaf and the peduncle of the ovule look alike; 
but transverse sections show two bundles in the petiole and four in 
the peduncle. Both cut easily in paraffin. 

Spermatogenesis. —The entire staminate cone, even at the time of 
shedding pollen, can be cut in paraffin. For the latest stages, how¬ 
ever, it is better to remove the sporophylls and cut them separately, 
since the sections must not be thicker than 5 /*, if they are to show 
the internal structures of the pollen grain. 

The young staminate cones become recognizable in June; by 
September, they have nearly or quite reached the spore mother¬ 
cell stage, but the division of the spore mother-cell does not 
take place until the following April. In these early stages the 
bud scales should be carefully dissected away before fixing. Pollen 
is shed early in May. Fix in chromo-acetic acid, with or without 
a little osmic acid, cut 5 n thick, and stain in iron-alum haema- 
toxylin. There are four cells in the pollen grain at the time of shed¬ 
ding (Fig. 97). 

Pollen tubes and their structures must be studied in sections of 
the nucellus. Fertilization, in the Chicago region, occurs about the 
middle of September. 

Oogenesis. —'Young ovules about 0.25 mm. in length are found 
about the middle of April; the megaspore mother-cell and its divi¬ 
sion into four megaspores are found about the first of May; the 
free nuclear stage in the development of the female gametophyte 
extends from the first week in May to the first week in July; during 
July, walls appear; then the archegonium initials and the growth of 
the archegonium, the ventral canal cell being cut off the second week 
in September; fertilization, free nuclear division in the sporophyte, 
and the beginning of walls may all be found before the end of Sep¬ 
tember; cotyledon stages belong to October, and when the seeds 
fall in November the embryo extends throughout nearly the entire 
length of the endosperm. This is the winter resting stage, but, 
planted in the greenhouse, the seeds germinate without any resting 
period, as in the case of cycads. 

For all stages in oogenesis and development of the embryo, use 
chromo-acetic acid. The free nuclear stages in both gametophyte 
and sporophyte are almost sure to plasmolyze. The special chromo- 
acetic-osmic-acid solution is better than one with less acetic acid. 
Hot alcoholic corrosive sublimate-acetic acid will cause very little 


278 


METHODS IN PLANT HISTOLOGY 



shrinking if the pieces are not too large; but figures are not as satis¬ 
factory as in chromic material. 

For sections of the entire ovule, use safranin, gentian-violet, 
orange; for free nuclear stages in both gametophyte and sporophyte, 


Fig. 97 —Ginkgo biloba: photomicrograph of microspores at shedding stage, showing two 
prothallial cells, generative cell, and tube cell. Stained in iron-alum haematoxylin. Eastman Com¬ 
mercial Ortho film, Wratten B filter (green); Spencer 4-mm. objective, N.A. .66; ocular X6, 
arc light; exposure, 3f seconds. Negative by Dr. P. J. Sedgwick. X654. 

use iron-haematoxylin with a touch of orange; for the megaspore 
membrane, safranin seems to be the best stain. 

GYMNOSPERMS—CONIFERALES 

Since Pinus is an available laboratory type, we shall describe 
methods for demonstrating various phases in the life-history of this 
genus, hoping that the directions will enable the student to experi¬ 
ment intelligently with similar forms. The dates are for Pinus 
Laricio in the vicinity of Chicago, but dates will be different for differ- 


SERPMATOPHYTES—GYMNOSPERMS 


279 


ent species and even for the same species in different regions; P. 
LariciOj at Chicago, sheds pollen about the middle of June, but P. 
maritima at Auckland, New Zealand, sheds its pollen about the 
first of October. After a year’s collecting in any region, there should 
be no difficulty, since the dates do not vary much from year to year. 

The Vegetative Structures.—The stem, root, and leaf will be 
treated separately. 

The stem .—The vascular cylinder is an endarch siphonostele, a 
type which, with few exceptions, is found throughout the living 
gymnosperms. 

The young stem in its first year’s growth is green and soft and is 
easily cut in paraffin. The best time to collect material is soon after 
the young shoot has emerged from the bud scales in the spring. 
Resinous material, like these young stems, do not fix well in aqueous 
media. With a thin safety-razor blade, cut the stem transversely 
into pieces about 5 mm. in length; fix in formalin alcohol, imbed in 
paraffin, and stain in safranin and light green. Longitudinal sections 
of the buds in winter or early spring condition are instructive for 
comparison with longitudinal sections of the ovulate cone. Trim 
away most of the bud scales and cut a slab from opposite sides, 
leaving a piece 2 or 3 mm. thick to be imbedded. The bud, and also 
selected pieces of the young stem, will show the structure of the young 
leaf. Later in the season, even the first year’s shoot should be cut 
without imbedding. The two- and three-year shoots and all older 
material can be cut freehand, without imbedding, and, preferably, 
before fixing. Such sections are transferred directly from the knife to 
95 per cent alcohol. 

For the structure of the adult stem, select a clear board and, for 
transverse sections, cut out pieces about 2 cm. long and 6 to 10 mm. 
square; for longitudinal sections, use pieces about 2 cm. long, with 
5 and 10 mm. for the other two faces. Cut from the face which will 
give sections 5X10 mm. Orient carefully, so that the longitudinal 
radial sections shall be exactly parallel with the rays, and the longi¬ 
tudinal tangential sections exactly tangential to the rays. Leave the 
sections in 95 per cent alcohol for 15 or 20 minutes before staining. 
Stain for at least 24 hours in safranin, extract the stain until only a 
faint red color is left in the cellulose walls, and then stain in Dela- 
field’s haematoxylin. Stain some of the sections in safranin and 
crystal-violet and some in safranin and light green. Of course, 


280 


METHODS IN PLANT HISTOLOGY 


every preparation should contain transverse, longitudinal radial, and 
longitudinal tangential sections. 

The safranin and Delafield’s haematoxylin should show the bars 
of Sanio very clearly, as should also the safranin and crystal-violet. 
If a preparation shows the bars of Sanio and differentiates the torus 
of the bordered pit, the technic is good (Fig. 98). 

Jeffrey’s maceration method will isolate the tracheids and other 
cells, which can then be stained and mounted in balsam. Such 
preparations show features which might be overlooked in sections. 

The root .—The primary root should be studied in the embryo 
while it is still contained in the seed ; Collect material in September, 
October, or at any later date. If material is collected in winter, the 
seeds should be soaked in water for a day or two before fixing. In 
any case, remove the testa and cut a thin slab from opposite sides 
of the endosperm to facilitate fixing and infiltration. For secondary 
roots and also for the structure of the stele in the primary root, 
germinate the seeds and fix material after the hypocotyl has reached 
a length of 3 or 4 cm. The seeds of Pinus edulis, commonly called 
Pinon, or edible pine, can be obtained in most cities. They are 
particularly good for a study of the mature embryo and the seedling. 

The older roots are treated like the stems. The structure of roots 
2 or 3 mm. in diameter is wonderfully regular (Fig. 99). 

The leaves .—The leaves of our common gymnosperms cut readily 
in paraffin while they are young and tender, but as they approach 
maturity it is a fuitless task to attempt paraffin sections. 

Good sections may be obtained in great quantities with little 
trouble by the following method: Make a bunch of the needles as 
large as one’s little finger, wrap them firmly together with a string, 
allowing about i inch of the bunch to project above the wrapping; 
dip two or three times into melted paraffin; and then dip into cold 
water to harden the paraffin. While there is no infiltration, the paraf¬ 
fin holds the needles in place for cutting. Fasten in a sliding micro¬ 
tome and cut with the knife placed obliquely. Place the sections 
in water as they are cut and the paraffin can be easily skimmed away. 
Then fix the sections in 95 per cent alcohol for half an hour; transfer 
to 70 per cent alcohol, where they should remain for about 5 minutes; 
then to 50 per cent alcohol, where they should be kept until the 
green color, due to chlorophyll, disappears. Stain in safranin and 
light green. 


SPERMATOPHYTES—GYMNOSPERMS 


281 


Spermatogenesis. —In October the clusters of staminate cones 
which are to shed their pollen in the coming spring are already quite 
conspicuous. The cones should be picked off separately, and the 



Fig. 98 .—Pinus Strobus: photomicrograph of longitudinal radial section of mature wood of 
stem, showing bordered pits and bars of Sanio. Safranin and crystal-violet, Eastman Commercial 
Ortho film, Wratten B filter (green); Spencer 4-mm. objective, N.A. .65; arc light; exposure, £ 
second. Negative by Dr. P. J. Sedgwick. X166. 













282 


METHODS IN PLANT HISTOLOGY 


scales should be carefully removed so as to expose the delicate 
greenish cone within. At this time the sporogenous cells are easily 
distinguished. Material collected in January, or at any time before 
growth is resumed in the spring, shows about the same stage of 
development. If it is desired to secure a series of stages with the 
least possible delay, a branch bearing numerous clusters of cones 
may be brought into the laboratory and placed in a jar of water. 
Growth is more satisfactory in case of branches broken off in the 



Fig. 99. —Picea nigra: photomicrograph of transverse section of root. Fixed in alcohol and 
stained in safranin and Delafield’s haematoxylin. Eastman Commercial Ortho film, Wratten B 
filter (green); J. Swift and Son 1-inch objective; arc light; exposure, l second. Negative by 
Dr. P. J. Sedgwick. X28. 

winter than in those brought in before there has been any period of 
rest. The material can be examined from time to time, and a com¬ 
plete series is easily secured. The mitotic figures in the pollen 
mother-cells furnish exceptionally instructive preparations. The 
two mitoses take place during the last week in April and the first 
week in May. Staminate cones which will yield mitotic figures can 
be selected with certainty by examining the fresh material. Crush 
a microsporangium from the top of the cone and one from the bottom, 
add a small drop of water and a cover to each, and examine. If 
there are pollen tetrads at the bottom, but only undivided spore 




SPERMATOPHYTES—GYMNOSPERMS 


283 


mother-cells at the top, it is very probable that longitudinal sections 
of the cone will yield the figures. If a drop of methyl green be 
allowed to run under the cover, it will enable one to see whether 
figures are present or not. When desirable cones are found, slabs 
should be cut from two sides, in order that the fixing agent may pene¬ 
trate more rapidly and that infiltration with paraffin may be more 
thorough. 

The later stages, showing the germination of the microspores, 
furnish better sections if the cones are cut transversely into small 
pieces about 5 mm. thick. It is very easy to get excellent mounts 
of the pollen just at the time of shedding, which, in Pinus Laricio 
in the vicinity of Chicago, occurs near the middle of June. Shake 
a large number of cones over a piece of paper, thus securing an 
abundance of material; slide the pollen off from the paper into a 
bottle half-full of water and shake a little to wet the pollen grains, 
because pollen of wind-pollinated plants is likely to be more or less 
shriveled at the time of shedding. A few minutes in water will 
make the pollen turgid. Fix in formalin-alcohol-acetic acid. If 
material is so abundant that you can lose much of it and still have 
plenty left, fix in chromo-acetic-osmic acid. Staining, especially 
the staining of mitotic figures, is likely to be more brilliant after 
fixing in the chromic series. However, most of the mitoses take 
place before the pollen is shed or after it reaches the nucellus. Infil¬ 
tration in the bath will not require more than 30 minutes. When 
the infiltration is complete, pour out into a paper tray or an imbedding 
dish, just warm enough to allow the pollen to settle to the bottom. 

A layer of pollen 3 mm. thick, with enough paraffin to make the cake 
about 5 mm. thick, will be satisfactory for cutting. Or, the paraffin 
may be poured out upon a piece of cold glass. Still another method 
is to leave the paraffin in a shell or vial during infiltration in the bath, 
and then let it cool in the bottle. After the paraffin is hard, break 
the bottle. Break the bottle carefully, cut off the lower portion of the ^ 
paraffin containing the pollen, mount it on a block in the usual man¬ 
ner, and trim away some of the paraffin so that two parallel surfaces 
will make the sections ribbon well. Sections should not be thicker 
than 5 m- Material in this stage shows a large tube nucleus, a some¬ 
what lenticular (generative) cell with a more deeply staining nucleus, 
and, lastly, two small prothallial cells quite close to the spore 
wall. The prothallial cells cannot always be detected at this stage, 


284 


METHODS IN PLANT HISTOLOGY 


and there may be some doubt as to whether two such cells are always 
present. The division of the lenticular cell into “stalk cell” and 
“body cell,” and also the division of the body cell into the two male 
cells, must be looked for in sections of the nucellus of the ovule. 

Abies balsamea is a better type for illustrating spermatogenesis, 
since the pollen mother-cells and the pollen grains are much larger 



Fig. 100. —Abies balsamea: photomicrograph of pollen; one complete section showing two 
prothallial cells, the stalk cell, generation cell, and tube cell with starch grains. Fixed and cut 
while still within the microsporangium. Stained in safranin and gentian-violet. Eastman Com¬ 
mercial Ortho film, Wratten B filter (green); Spencer 4-mm. objective, N. A. .66; ocular X6; arc light; 
exposure, 6 seconds. Preparation by Dr. A. H. Hutchinson, negative by Dr. P. J. Sedgwick. X 784. 


and the division of the generative cell into the “stalk” and “body” 
cells takes place before the pollen is shed (Fig. 100). 

Araucaria and Agathis are the best forms for illustrating numerous 
prothallial cells/ Podocarpus and Taxodium are also good. Thuja or 
Juniperus may be used to illustrate the entire absence of prothallial 
cells, a very advanced condition; while both these genera have highly 
developed sperms, like those of the cycads and Ginkgo, except that 
they lack cilia. This is a good illustration of the fact that one 
structure may advance while another remains primitive. 





SPERMATOPHYTES—GYMNOSPERMS 


285 


Oogenesis.—In Pinus Laricio the rudiment of the ovulate strobi- 
lus, which is to be pollinated in June, can be detected in the preceding 
October. The collection of this stage is very uncertain, because there 
seems to be no mark distinguishing buds containing ovules from buds 
which are only vegetative. By collecting numerous buds from the 
tops of vigorous trees which are known to produce an abundance of 
strobili, a few buds containing the desired stages may be obtained. 
In May, after the strobili break through the bud scales, material 
is easily collected. Up to the time of pollination the entire ovulate 
strobilus cuts easily in paraffin. Longitudinal sections of the cone at 
this time give good views of the bract and ovuliferous scale bearing 
the ovules. The integument is very well marked, and in the nucellus 
one or more sporogenous cells can usually be distinguished. As 
soon as the scales close up after pollination, the cone begins to harden 
and soon makes trouble in cutting. Even before the scales close up, it 
is better to cut a slab from opposite sides of the cone; after the scales 
close, it is almost a necessity. For sections of the whole cone, fix 
in formalin alcohol. Dr. Hannah Aase succeeded in cutting complete 
series of paraffin sections from cones of Pinus Banksiana more than 
2 cm. in length. She fixed them in formalin alcohol, and used pro¬ 
longed periods in dehydrating, clearing, and infiltrating. Land’s 
bichromate of potash and glue fixative was used in fixing the sections 
to the slide. Such series of sections of large cones were necessary for 
an investigation of the vascular anatomy. 

For a study of the ovule and the structures within it, better 
preparations will be obtained by carefully cutting off the pair of 
ovules from the scale. In free nuclear stages of the female gameto- 
phyte, which begin in the autumn, are interrupted by winter, and are 
completed in May, plasmolysis is likely to occur. After walls appear 
there is less danger. From the middle of May to the first of July 
collections should be made at intervals of two or three days, since 
during these six weeks the gametophyte completes the free nuclear 
stage and develops cell walls, the archegonium completes its entire 
development, the egg is fertilized, and the sporophyte may reach the 
suspensor stage. 

At the stages shown in Figure 101 B-D, it is a good plan to remove 
the female gametophyte with its proembryos from the ovule; but 
at the stages shown in Figures 10L4 and 102 the pollen tubes with 
their contents are rapidly working their way through the nucellus 


286 


METHODS IN PLANT HISTOLOGY 


toward the archegonia, and consequently, in some of the material, 
it is better to retain enough of the tissues of the ovule to keep the 
nucellus in place. In later stages, after fertilization has taken place, 
the developing testa should be removed with great care, for a very 
slight pressure is sufficient to injure the delicate parts within. 

With any fixing agent of the chromic-acid series, the free nuclear 
stages of the female gametophyte and, later, the archegonia and 
proembryo stages like those shown in Figure 101, are likely to show 



Fig. 101. Pinus Laricio: A, top of prothallium with an archegonium just before the cutting 
off of the ventral canal cell; fixed in Flemming’s weaker solution and stained in Haidenhain’s iron- 
alum haematoxylin; collected June 18, 1897; B, C, and D, early stages in the development of the 
embryo; fixed in chromo-acetic acid and stained in safranin, gentian-violet, orange- collected 
July 2, 1897. X104. 

plasmolysis. Stages like those shown in Figure 101 B-D, without 
any shrinking, were secured by Miss Ethel Thomas by using hot 
alcoholic corrosive sublimate-acetic acid with formalin (4 g. corrosive 
sublimate, 5 c.c. acetic acid, 5 c.c. formalin, 100 c.c. 70 per cent alcohol). 
Figures like that shown in Figure 101 are better in chromo-acetic- 
osmic acid, but, in general, we should recommend the corrosive 
sublimate solution. 

The period at which the various stages may be found varies 
with the species, the locality, and the season. In Pinus Laricio 


SPERMATOPHYTES—GYMNOSPERMS 


287 



Fig. 102 .—Pinus Laricio: photomicrograph showing the formation of the ventral canal cell; 
usually this cell is not so large in proportion to the egg; fixed in Flemming’s weaker solution and 
stained in safranin, gentian-violet, orange; the preparation was made in 1897, the photomicrograph 
in 1915; Cramer contrast plate; 4-mm. objective; ocular X4; Abb§ condenser; yellowish-green 
filter and also a strong filter used in outdoor photography; camera bellows, 75 cm.; arc light; 
exposure, 7 minutes. Negative by Miss Ethel Thomas. X587. 

and sperm nuclei occurred about a week later, and stages like Figure 
1015, C, and D, were common in material collected July 2. In the 
season 1896 all the stages appeared about 2 weeks earlier. In Pinus 
sylvestris the stages appeared a little earlier than in Pinus Laricio. 


the megaspore mother-cells appear as soon as the young strobili 
break through the bud scales. At Chicago, in the season of 1897, 
material collected May 27 did not yet show archegonia; the ventral- 
canal cell was cut off about June 21 (see Fig. 102), the fusion of the egg 




288 


METHODS IN PLANT HISTOLOGY 



After the stage shown in Figure 101A has appeared, it is necessary 
to collect every day until the stage shown in Figure 101D is reached. 
If collections are made at intervals of 3 or 4 days, the most interesting 

stages, like the cutting off 
of the ventral-canal cell, 
fertilization, and the first 
divisions of the nucleus of 
the oospore, may be missed 
altogether. It should be. 
mentioned that all the 
ovules of a cone will be in 
very nearly the same stage 
of development; conse¬ 
quently, it is worth while 
to keep the ovules from 
each cone separate. 
Stages like that shown in 
Figure 102 are rare in mis¬ 
cellaneous collections, but 
if ovules from each cone 
are kept separate and this 
figure is found, the rest 
of the ovules from that 
cone will be likely to show 
some phase of this interest¬ 
ing mitosis. 

Thuja and Juniperus 
are good types to illustrate 
the archegonium complex 
and the large, highly or¬ 
ganized male cells. In 
Thuja a series from the 
appearance of archego¬ 
nium initials to young 
embryos may be collected between June 10 and June 20. In 
Juniperus pollination occurs late in May and fertilization takes place 
12J months later. The megaspores are formed late in April and 


Fig. 103 .—Pinus Banksiana: photomicrograph of young 
embryos teased out by the method described in the text; 
from a preparation by Dr. J. T. Buchholz; Cramer contrast 
plate; 16-mm. objective; no ocular or Abb6 condenser; 
camera bellows, 75 cm.; safranin filter; arc light; exposure, 
17 seconds. Negative by Miss Ethel Thomas. X54. 


the development of the 
weeks. 


female gametophyte occupies about 6 




SPERMATOPHYTES—GYMNOSPERMS 


289 


The Embryo.—The early stages of the sporophyte, usually 
designated as the proembryo, have been mentioned already. 

From the time when the suspensors begin to elongate up to the 
appearance of cotyledons, instructive preparations can be made by 
mounting the embryo whole. Dr. J. T. Buchholz has developed a 
method for handling these small objects. Remove the testa and 
then, under water, hold the endosperm gently with forceps and 
press the neck and upper part of the archegonium with a needle, 
pressing, and at the same time drawing the needle away, so as to 
pull the young embryo out. Some of the embryos will be broken, but 
by careful manipulation more than half should be entirely uninjured. 
Fix in formalin (5 per cent in water), stain in Delafield’s haematoxylin, 
transfer to 10 per cent glycerin, and continue with the Venetian 
turpentine method. A preparation made in this way is shown in 
the photomicrogaph (Fig. 103). 

These stages, and all subsequent stages, are easily cut in paraffin 
without removing the embryo from the endosperm. Cut a thin 
slab from opposite sides of the endosperm, fix in chromo-acetic acid, 
with or without a little osmic acid, imbed in paraffin, and stain in 
safraqin and gentian-violet. This will give a good view of the abun¬ 
dant starch and other food stuff, «,nd at the same time will bring out 
sharply the cell walls of the embryo. 

GYMNOSPERMS—GNETALES 

Of the three peculiar genera belonging to this order only one, 
Ephedra, occurs in the United States. Welwitschia is found only in 
Damaraland, South Africa, and Gnetum is tropical and subtropical. 
Gnetum thrives in the greenhouse, but the other two have not been 
cultivated successfully. All show vessels in the secondary wood, an 
angiosperm character. The strobili can be fixed in formalin-alcohol- 
acetic acid, but in Ephedra the dry chaffy scales must be dissected 
away before completing the dehydration and infiltration with paraffin. 
If you can secure material of Ephedra, Dr. Land’s researches 1 pre¬ 
sent a very complete life-history, with dates of various stages and 
suggestions for fixing and staining. 

1 Botanical Gazette , 38:1—18, 1904, and 44:273—292, 1907. 


CHAPTER XXV 


SPERMATOPHYTES 
A NGIO SPERM S 

If one should master the technic for all features of this group, he 
would be prepared to deal with all the rest; for embryo sacs are so 
delicate that they are as difficult as the free nuclear stage in the female 
gametophyte of a gymnosperm, while the peach stone needs a petro- 
tome rather than a microtome. Between these extremes there is 
everything imaginable in structure, chemical composition, and 
consistency. 

Some hints will be given, but the student will gradually learn 
what should be cut freehand and what should be imbedded; what 
stages in floral development, what stages in the development of the 
embryo sac, or what stages in spermatogenesis are likely to be corre¬ 
lated with easily recognized field characters, and what fixing agents 
are likely to give the best results, with various kinds of material. 

The Vegetative Structures—In stems, roots, and leaves the 
more delicate structures should be imbedded in paraffin and the more 
rigid structures should be cut without imbedding at all; but it should 
be remembered that the range of structures which can be imbedded 
in paraffin can be increased by the use of hydrofluoric acid; and also 
that hydrofluoric acid or cellulose acetate will extend almost indefi¬ 
nitely the range of structures which can be cut without imbedding. 

The stem .—The vascular cylinder of the angiosperms is either an 
endarch siphonostele, or a polystele derived from it. For a study of 
the development of the stem, the common geranium ( Pelargonium ) 
may be recommended. Near the base of a fresh stem, about 1 cm. 
in diameter, cut freehand sections and fix them in 95 per cent alcohol 
for 10 to 20 minutes; transfer to 70 per cent alcohol to extract the 
chlorophyll, and then stain in safranin and light green. Such sections 
will show both primary and secondary structures in the stele and in 
the cortex. Higher up, there will be secondary structures only in 
the stele; and still higher up will be found the origin of interfascicular 
cambium. All these can be cut without imbedding, but the earlier 

290 


SPERMATOPHYTES—ANGIOSPERMS 


291 


stages showing the differentiation of protoxylem, metaxylem, and 
the origin of secondary xylem are too soft for successful freehand 
sections. Fix in alcohol-formalin-acetic acid (10 c.c. formalin, 
5 c.c. acetic acid, and 100 c.c. of 50 per cent alcohol) and imbed in 
paraffin. 

For a study of woody stems, Tilia americana (basswood) is good, 
and shoots from 5 to 10 mm. in diameter are easy to cut. Very hard 
stems like Hicoria (hickory) and Quercus (oak) must be boiled and 
treated with hydrofluoric acid, if you expect to cut shoots more than 
5 to 7 mm. in diameter. However, with a good sharp knife and a 
rigid microtome much larger sections can be cut without resorting 
to hydrofluoric acid. Of course, veneer machines cut very large and 
fairly thin, smooth sections from the most refractory woods. 

While a random selection of stems would furnish material for 
practice in technic, we suggest that the stem of Clintonia shows a good 
siphonostele in a monocotyl; the rhizome of Acorus calamus is a 
good type for the amphivasal bundle and, although a monocotyl 
still shows a differentiation into stele and cortex; Zea Mais , univer¬ 
sally used but not characteristic of monocotyls, shows scattered 
bundles, but not the amphivasal condition; Aloe , Dracaena , or 
Yucca will illustrate secondary wood in monocotyls. Iris has a 
highly developed endodermis in the rhizome; and Nymphea or Nuphar 
will show scattered bundles in a dicotyl. 

Lenticels and tyloses are abundant and typical in Menispermum , 
and very thin sections can be cut without imbedding; but both 
these structures are well developed while the stem can still be cut 
in paraffin without previous treatment in hydrofluoric acid. 

The sieve tubes of the phloem are easily demonstrated in Cucurbita 
Pepo, the common pumpkin; other members of the family furnish 
good material. Take pieces of stem about 1 cm. long and not too 
hard to cut in paraffin, fix in formalin alcohol, and stain in safranin, 
gentian-violet, orange. The tropical Tetracera, one of the Dillenia- 
ceae, has sieve plates so large that they are easily seen with a pocket 
lens. The phloem area is so large in the larger stems that it can be 
cut out for imbedding in paraffin long after the entire stem has become 
too hard for paraffin sections. Safranin and gentian-violet is a good 
stain for sieve tubes. It was once thought that these large sieve 
tubes afforded an obvious illustration of the continuity of protoplasm; 
but, as a matter of fact, the actual protoplasmic connections are 


292 


METHODS IN PLANT HISTOLOGY 


scanty and hard to demonstrate. Iron-alum haematoxylin and 
orange will differentiate the strands if you are careful. 

Roots. —It has long been known that the root-tip furnishes- 
constantly available material for a study of mitosis (Fig. 104). An 
onion thrown into a pan of water will soon send out numerous roots. 
Soak beans in water for several hours and then plant them in loose, 
moist sawdust. In a greenhouse, with “ bottom heat,” the primary 
root will be long enough in 2 or 3 days. The large, flat beans, espe¬ 
cially Vida Faba, are very favorable. The root-tips of Trillium 
grandiflorum , Tradescantia virginica, Podophyllum peltatum, Arisaema 
triphyllum, and Cypripedium pubescens may be mentioned as known 
to be favorable; but it is very possible that the best root-tip has not 
yet been tried. 

Cell division does not proceed with equal rapidity at all hours 
of the day. Kellicott has shown that in the root-tips of Allium 
there are in each 24 hours two periods at which cell division is at 
the maximum, and two at which it is at the minimum. The maximum 
periods are shortly before midnight (11:00 p.m.), and shortly after 
noon (1:00 p.m.). The minima, when cell division is at the lowest 
ebb, occur about 7:00 a.m. and 3:00 p.m. When cell division is most 
vigorous, there is little elongation, and when cell division is at the 
minimum, cell elongation is at the maximum. Consequently, 
root-tips of Allium should be collected about, 1:00 p.m. or 11:00 p.m. 
Lutman, later, made observations upon periodicity of mitosis in the 
desmid, Closterium; and in 1915, Karsten made a comparatively 
extended study of periodicity in various stems and roots, together 
with notes on algae. 

It is safe to say that the maximum number of mitoses in root-tips 
will be found shortly after noon (1:00 p.m.) and shortly before mid¬ 
night (11:00 p.m.) (Fig. 104). It is certain, however, that abundant 
mitoses may be found at other times—even at 3:00 p.m. —in sporangia 
of ferns, in anthers of angiosperms, in endosperm, and in free nuclear 
stages of the embryo of gymnosperms. 

Mitotic figures play such an important part in the development 
of the plant and in modern theories of heredity that it is worth while 
to acquire a critical technic in fixing and staining these structures. 
Use the various fixing agents—Flemming’s weaker solution, chromo- 
acetic acid with or without a little osmic, Benda’s fluid, Bouin’s 
fluid, corrosive sublimate with acetic acid, and any others. Make 


SPERM ATOPHYTES—ANGIOSPERMS 


293 



Fig. 104.— Trillium grandiflorum: photomicrograph of longitudinal section of root-tip, 
fixed in chromo-acetic-osrnic acid at 1:00 p.m., and stained in iron-alum haematoxylin. Eastman 
Commercial Ortho film, Wratten H filter (blue); Spencer 2-mm. fluorite objective; ocular X6; 
arc light; exposure, 12 seconds. Negative by Dr. P. J. Sedgwick. X 1,174. 



294 


METHODS IN PLANT HISTOLOGY 


yourself master of Haidenhain’s iron-alum haematoxylin; then add 
the safranin, gentian-violet, orange combination; then safranin and 
anilin blue; and then experiment for yourself, but remember that the 
triumphs of modern cytology have been won with iron-haematoxylin 
and that you cannot read intelligently the literature of the past 
twenty-five years until you have gained at least an approximate 
mastery of this stain. Of course, dehydration, clearing, and infiltra¬ 
tion must be very gradual. The schedules by Yamanouchi and by 
Sharp, on pages 45 and 46, will repay careful study. 

In staining with safranin, gentian-violet, orange, allow the 
alcoholic safranin to act for 16 to 24 hours; then extract it with 50 
per cent alcohol, slightly acidulated with hydrochloric acid, if neces- 
sary, until the stain has almost disappeared from the spindle; then 
pass through 70, 85, 95, and 100 per cent alcohol; stain in gentian- 
violet dissolved in clove oil, or in a mixture of clove oil and absolute 
alcohol, for 5 to 20 minutes; follow with orange dissolved in clove 
oil, remembering that this will weaken the safranin and sometimes 
the gentian-violet; finally use pure clove oil to differentiate the 
gentian-violet. Leave the slide in xylol for 2 to 5 minutes to remove 
the clove oil and to hasten the hardening of the balsam. 

If you use aqueous gentian-violet or crystal-violet, after the 
safranin is satisfactory, transfer to water and then to the violet. 
After staining in violet, dip in water to remove the excess of stain 
and then dehydrate rapidly in 95 per cent and absolute alcohol, 
differentiate in clove oil, and then transfer to xylol. 

The structure and development of the young root will be shown, 
to some extent, in preparations made for mitotic figures. The origin 
of dermatogen, periblem, plerome, and also of protoxylem, is well 
shown in Zea Mays. An ear of sweet corn, as young and tender as 
you can find on the market, will furnish material. Cut out from the 
grain a rectangular piece about 2X3 mm. and 4 or 5 mm. long; if 
you want to show also the structure of the entire grain, take a section 
the entire length of the grain, perpendicular to the flat side of the 
grain, and about 2 mm. wide. Cut the latter longitudinally; the 
rectangular pieces are sufficient for transverse sections. Fix in 
chromo-acetic acid. The roots of Hordeum vulgare (barley) might 
also be suggested. 

The roots of Ranunculus repens and Sambucus nigra furnish 
good illustrations of the radial arrangement of xylem and phloem. 


SPERMATOPHYTES—ANGIOSPERMS 


295 


Smilax shows the radial arrangement, with a large number of poles 
and a very highly differentiated endodermis. The origin of secondary 
xylem and phloem is well shown in Sambucus nigra. Vida Faba 
shows very clearly the origin of secondary roots. Pislia stratiotes, 
although not so generally available, is splendid for showing the 
origin of secondary roots. The arrangement of cells in the young 



Fig. 105 .—Sparganium eurycarpum: photomicrograph of transverse section of young root; 
fixed in chromo-acetic acid and stained in Bismarck brown; Cramer contrast plate; 16-mm. 
objective; ocular X4; no Abb6 condenser; yellowish-green filter; camera bellows, 1 meter; 
exposure, 8 seconds. X90. 

roots of aquatic or semi-aquatic plants is often extremely regular 
(Fig. 105). 

The leaf .—Young and tender leaves should be fixed in formalin 
alcohol and cut in paraffin. Cut sections freehand whenever there 
is sufficient rigidity. Resort to pith only when necessary. In 
cutting sections of a leaf like that of Lilium , lay one leaf on another 
until you have a bundle of them which will be nearly square in trans¬ 
verse section. Wrap the bundle with string for about 15 mm.; cut 



296 


METHODS IN PLANT HISTOLOGY 


the bundle transversely so that about 5 mm. of the bundle will 
project beyond the tied portion. Dip in melted paraffin, as already 
suggested for Pinus, fasten the tied portion in the sliding microtome, 
and cut with the knife placed obliquely. About 15 to 20 p is a good 
thickness for general leaf structure. In case of large leaves, cut 1 cm. 
wide and 3 cm. long and tie them together to make a good bundle for 
cutting. 

Of course, for the finest preparations, imbed in paraffin. The 
common lilac, Syringa, has a good leaf to illustrate palisade and spongy 
parenchyma; the privet, Ligustrum , is also excellent. 

Buds will furnish beautiful preparations of young leaves and, at 
the same time, will show the vernation. Cut the bud transversely, 
a little above the middle; remove the bud scales, if they promise 
to cause trouble; retain only enough tissue at the base of the bud 
to hold the parts in place. Fix in formalin alcohol and stain in 
safranin and light green. 

Epidermis stripped from the leaf, fixed in 10 per cent formalin 
in water for a day or two, and then stained in safranin and anilin 
blue, will give excellent views of stomata. The development of 
stomata is particularly well shown in Sedum purpurascens, even in 
leaves which have reached the adult size. The epidermis is very 
easily stripped from a leaf of Sedum. If the big Sedum maximum is 
available, pieces of epidermis 6 or 7 cm. long and 2 or 3 cm. wide are 
easily stripped off, almost free from any underlying tissue. The 
epidermis of Lilium and Tradescantia show fine, large stomata, but 
it is not so easy to strip off. In these two genera the stomata, as in 
nearly all leaves, show only the adult structure. 

Floral Development.—For a study of floral development very 
young buds are necessary, and it is best to select those forms which 
have rather dense clusters of flowers, in order that a complete series 
may be obtained with as little trouble as possible. 

The usual order of appearance of floral parts is (1) calyx, (2) 
corolla, (3) stamens, and (4) carpels; but if any of these organs is 
reduced or metamorphosed, their order of appearance may be affected. 

Floral development is easily studied in the common Capsella 
bursa-pastor is. The best time to collect material is late in March 
or early in April. Dig up the plant, carefully remove the leaves, 
and in the center of the rosette a tiny white axis will be found. A series 
of these axes from 3 to 9 mm. in length, and from 1.5 to 3.5 mm. 


SPERMATOPHYTES—ANGIOSPERMS 


297 


in diameter will give a very complete series of stages in the 
development of the floral organs. Preparations from the apex 
of the shoot taken after the inflorescence appears above ground are 
not to be compared with those taken early in the season, because 
the pedicels begin to diverge so early that median longitudinal 
sections of the flowers are comparatively rare. Fix in chromo-acetic 
acid and stain in Delafield’s haematoxylin. The sections should be 
longitudinal and about 5 p thick. Capsella shows the hypogynous 
type of development. The order of appearance of floral parts is 
(1) calyx, (2) stamens, (3) carpels, and (4) petals. The ovary is 
compound (syncarpous). 

Ranunculus, which is also hypogynous, will illustrate the develop¬ 
ment of the simple (apocarpous) ovary. The ovules appear quite 
early, so that the archesporial cell, or even the megaspores, may be 
seen while the carpel is still as open as in any gymnosperm. The 
whole structure is a simple strobilus. 

Rumex crispus (Yellow Dock) is also a good hypogynous type, and 
the densely clustered flowers afford a fine series of stages. Besides, 
in transverse sections, the early stages in spermatogenesis are very 
clear. 

In the willows, Salix , the bud scales must be removed and the 
copious hairs should be trimmed off as much as possible with scissors, 
after which the catkin should be slabbed a little on opposite sides 
to facilitate penetration. This is a fine illustration of a compound 
strobilus. 

The cat-tail, Typha, presents a simple type of floral development. 
The leaves should be dissected away long before the flowers can be 
seen from the outside. The cylindrical clusters, varying in diameter 
from 2 or 3 mm. up to the size of one’s finger, will afford a complete 
series of stages. Until the spadix reaches the diameter of a lead 
pencil, transverse sections are easily cut. For later stages, the outer 
part of the spadix should be sliced off so that only enough spadix is 
retained to hold the florets in place. 

Prunus and many other members of the Rosaceae furnish examples 
of the perigynous type of development. In many of them the floral 
parts do not occur in the usual succession. 

The epigynous type is well shown in the Compositae. The order 
of appearance is (1) corolla, (2) stamens, (3) carpels, and (4) calyx 
(pappus). 


298 


METHODS IN PLANT HISTOLOGY 


The common dandelion, Taraxacum officinale, affords an excellent 
series with little labor. Examine vigorous plants which have, as yet, 
no flowers or buds in sight. Dig up the plant and dissect away the 
leaves. If there is a white cluster of flower buds, the largest not 
more than 4 mm. in diameter, cut out the cluster, leaving only enough 
tissue at the base to hold the buds in place. Larger heads should be 
cut separately. 

Our most common thistle, Cirsium lanceolatum, shows the floral 
development with unusual clearness, but the preparation of the 
material is somewhat tedious. The involucre, which is too hard to 
cut, must be carefully dissected away. Retain only enough of the 
receptacle to hold the developing florets in place. A series of sizes 
with disks varying from 3 to 10 mm. in diameter will show the 
development from the undifferentiated papilla up to the appearance 
of the archesporial cell in the nucellus of the ovule. The Canada 
thistle, Cirsium arvense, is equally good, but it is more difficult to 
dissect out the desirable parts. In the common sunflower, Helian- 
thus annuus, the young floral parts, like the mature head, are so very 
large that a satisfactory study may be made with a low-power objec¬ 
tive. As in the case of the thistle, the involucre must be trimmed away 
and only enough of the receptacle retained to hold the florets together. 

Erigeron (we have cut E. philadelphicus and E. annuus) furnishes 
a beautiful example of epigynous floral development, and the heads 
are so densely clustered that, in a single section, one may find various 
stages from heads with undifferentiated disk up to heads with florets 
showing pappus, corolla, stamens, and carpels. 

Spermatogenesis.—The earlier stages in spermatogenesis will 
be found in the preparations of floral development. The origin of 
the archesporium, the origin of sporogenous tissue, and the formation 
of the tapetum are beautifully shown in longitudinal and in transverse 
sections of the anthers of Taraxacum and many other Compositae. 
Transverse sections of the head of Taraxacum or any similar head 
at the time when pollen mother-cells are rounding off in the center of 
the head, will show various stages from the mother-cells in the center 
to the tetrads of spores at the periphery. Transverse sections of 
the anther of Polygala give exceptionally well-defined views of the 
archesporial cells and sporogenous areas. 

Lilium, Trillium, Galtonia, Iris, Tradescantia, Vida, and Podo¬ 
phyllum can be recommended for demonstrating the nuclear changes 


SPERMATOPHYTES—ANGIOSPERMS 


299 


involved in the formation of spores from the mother-cell (Fig. 106). 
Several species of Lilium are common in greenhouses, and these 
may be used where wild material is not available. In early stages, 
where the sporogenous cells have not yet begun to round off into spore 
mother-cells, it is sufficient to remove the perianth, retaining just 
enough of the receptacle to hold the stamens in place. Transverse 
sections show the six stamens and also the young ovary. After the 



Fig. 106 .—Lilium candidum: photomicrograph of mitosis in pollen mother-cells; in one of 
the pollen mother-cells the twelve chromosomes can be counted; from a preparation by F. L. 
Pickett. Negative by Miss Ethel Thomas. X260. 

spore mother-cells have begun to round off,- each stamen should be 
removed so as to be cut separately. In securing the desirable stages 
showing the division of the mother-cell into microspores, much time 
and patience will be saved by determining the stage of development 
before fixing the material. Mitosis is more or less simultaneous 
throughout an anther. Long anthers are particularly favorable, 
since they may show a very closely graded series of the various 
phases of mitosis. An anther of Lilium may show mother-cells with 
nuclei in synapsis at the top, while the mother-cells at the bottom 
have reached the equatorial plate stage of the first division; or, the 






300 


METHODS IN PLANT HISTOLOGY 


mother-cells at the top may show the first division, while those at 
the bottom show the second. Determine the stage by examining a 
few mother-cells before fixing. 

From what has been said, it is evident that longitudinal sections 
should be cut to show mitosis. Transverse sections should be cut 
to show the general structure of the anther. It is not necessary to 
cut the stamens into pieces before fixing, since they are easily pene¬ 
trated and infiltrated; in later stages the stamens must not be cut 
into pieces, since the pollen grains and even the pollen mother-cells 
are easily washed out. 

The problem of fixing spore mother-cells has received much atten¬ 
tion. In fixing mother-cells and the two mitoses by which spores 
are formed, investigators have used almost exclusively the chromo- 
osmo-acetic acid solutions of Flemming, some preferring the weaker 
solution and some the stronger. These solutions have been used 
in nearly all of the work of the Bonn (Germany) school. Osterhout 
experimented with forty fixing agents, and then concluded that 
Flemming’s stronger solution was the best. Professor Gregoire and 
his students have made this their principal fixing agent. 

In the third edition of this book we stated that, in spite of the 
weight of authority, we believed the value of solutions with such 
a large proportion of osmic acid had been overestimated and we sug¬ 
gested that half the amount of osmic recommended in Flemming’s 
weaker solution would be sufficient. The weaker solution has 10 c.c. 
of 1 per cent osmic acid in 100 c.c. of the fluid. After an examination 
of thousands of figures in both reduction and vegetative divisions, 
we feel sure that 5 c.c. of the osmic is enough and, perhaps, 3 or 4 c.c. 
would be better. The formula for Flemming’s weaker solution is 
often given as follows: 


f Chromic acid (1 per cent). 25 c.c. 

A \ Glacial acetic acid (1 per cent). 10 c.c. 

(Water. 55 c.c. 

B. Osmic acid (1 per cent)_;. io c.c. 


Keep the mixture A made up, and add B as the reagent is needed 
for use, since the solution does not keep well. One seldom uses this 
reagent in large quantities. About 40 c.c. is as much as one is likely 
to need for any collection of anthers or root-tips. Take 36 c.c. of A 
and 4 c.c. of B. It will be worth while to try 36 c.c. of A and 2 c.c. 






SPERMATOPHYTES—ANGIOSPERMS 


301 


of B, or even 1 c.c. of B. If the regular formula is used, we should 
let it act for an hour, and then replace it by A, without any osmic 
acid. The osmic acid undoubtedly accelerates the killing of the 
protoplasm. This is seen more readily in animals. If Cycloys be 
brought into 30 c.c. of the solution A, the animals will swim for awhile; 
if 5 or 6 drops of 1 per cent osmic acid be added to the solution, the 
animals cease their movements almost instantly. Doubtless the 
osmic acid has the same effect upon plant protoplasm. Where 
fixing is slow, very few mitotic figures are found with the chromo¬ 
somes midway between the equator and the poles. The addition of 
10 drops of 1 per cent osmic acid to 50 c.c. of the solution just men¬ 
tioned will secure as large a proportion of anaphases as solutions 
which are stronger in osmic acid, and there is no disagreeable 
blackening. 

Farmer and Shove, in studying these mitoses and also vegeta¬ 
tive mitoses in Tradescantia , secured better results with a mixture 
of 2 parts of absolute alcohol and 1 part glacial acetic acid. They 
allowed the fixing agent to act 15 to 20 minutes, then washed in 
absolute alcohol, and imbedded by the usual methods. This propor¬ 
tion of acetic acid seems entirely too large for any accurate work 
with chromatin, and we doubt whether the structure of the cytoplasm 
is normal when so much acetic acid is used. 

The entire pollen mother-cell may be stained and mounted without 
sectioning. Two descriptions of technic appeared in 1912, one by 
Mann and the other by Pickett. Mann removes the pollen mother- 
cells before fixing and staining; Pickett fixes and stains the anther 
in toto and teases out the pollen mother-cells just before mounting. 

In Mann’s method, the anther is placed in a drop of water and 
the tip is cut off; a gentle tapping with a needle will then cause the 
pollen mother-cells to float out into the drop. Fix in Bouin’s fluid, 
4 to 8 hours, wash in 50 per cent alcohol until no color remains, and 
then stain in iron-haematoxylin. At this stage we should put the 
material into 10 per cent glycerin and follow the Venetian turpentine 
method. 

Pickett fixed entire anthers in chromo-acetic acid for 30 hours, 
washed in water for 24 hours, and then passed up to 80 per cent alco¬ 
hol. At this point, he stained in strong cochineal or Kleinenberg’s 
haematoxylin for 5 days, then completed the dehydration, cleared in 
cedar oil, teased out the mother-cells, and mounted in balsam. 


302 


METHODS IN PLANT HISTOLOGY 



In dealing with the whole anther, it is necessary to select stains 
which will not overstain. Alum cochineal and Mayer’s haem-alum 
might be suggested. It would be worth while to try a combination 
of the two methods. Fix the entire anther in chromo-acetic acid, 

wash in water, and then stain 
in iron-haematoxylin. When 
the last stage in staining is 
reached—the extraction of the 
stain in iron-alum—remove the 
pollen mother-cells and watch 
the differentiation; then wash 
in water and follow the Vene¬ 
tian turpentine method. 

The pollen grain at the time 
of shedding generally consists 
of two cells, the tube cell and 
the generative cell, which 
afterward divides and forms 
two male cells or two male 
nuclei. Lilium and Erythro- 
iiium furnish good illustrations 
of pollen shed in the two-cell 
stage (Fig. 107). In Silphium, 
Sambucus, and Sagittaria the 
generative nucleus divides 
before the pollen is shed. The 
division of the generative cell 
to form the two sperms takes 
place just before fertilization; 
consequently, in forms like 
Silphium, fertilization is likely 
to occur within less than 72 hours after the division of the genera¬ 
tive cell. 

Sections should not be more than 5 ju thick, if they are to show 
a clear differentiation of exine, intine, starch, and other structures. 
If sections have been stained in iron-haematoxylin, staining in safranin 
for from 3 to 7 minutes will give the exine a bright-red color and will 
not obscure the haematoxylin. A rather sharp stain in gentian- 
violet will stain the starch and also the intine. In Asclepias and many 


Fig. 107 .—Erythronium americanum: photomi¬ 
crograph of mature pollen grains; the one at the 
top, which is cut longitudinally, shows both the 
tube nucleus and the conspicuous generative cell; 
the other is cut transversely and shows the genera¬ 
tive cell, but not the tube nucleus; stained in 
safranin and gentian-violet; from a preparation by 
Dr. Lula Pace; Cramer contrast plate; 4-mm. 
objective; ocular X4; yellowish-green filter; bel¬ 
lows, 85 cm.; exposure, 3 minutes. X615. 


SPERMATOPHYTES—ANGIOSPERMS 


303 


orchids, in which a common exine surrounds the entire mass of pollen 
grains, care must be taken not to overstain. 

In many cases the pollen grains will put out their tubes in a 2 to 
5 per cent solution of cane-sugar in water. Where the interval 
between pollination and fertilization is known (about 72 hours in 
Lilium philadelphicum and 96 to 100 hours in L. canadense), pieces 
of the stigma and style showing pollen tubes can be selected with 
some certainty. 

Oogenesis.—As in spermatogenesis, the early stages will be found 
in preparations of floral development. The preparations of Capsella 
will show the origin and development of the nucellus (megaspo¬ 
rangium) and also the megaspore mother-cell. The division of the 
megaspore mother-cell to form four megaspores takes place shortly 
before the bud begins to unfold. A massive megasporangium with 
several megaspore mother-cells may be found in Ranunculus; a 
megasporangium with only one megaspore mother-cell and only one 
layer of cells surrounding it may be found in any of the Compositae. 
Senecio aureus and Erectites hieracifolium are good and are particu¬ 
larly easy to cut. In Trillium and in Cypripedium the embryo sac is 
formed from two megaspores, which are not separated by walls. In 
Lilium , Tulipa, Fritillaria, Erythronium, and many others, the embryo 
sac is formed by alb four megaspores, which are not separated by 
walls. In Peperomia, the Peneaceae, and some species of Euphorbia, 
the sac is formed by the four megaspores, not separated by walls, and 
the sac has 16 free nuclei. In Plumbagella the four megaspores, 
not separated by walls, constitute the mature sac, one of the mega¬ 
spores functioning as the egg, two more fusing to form the endosperm 
nucleus, while the fourth megaspore aborts; so that the embryo sac, 
ready for fertilization, contains only two nuclei. 

The reduction of chromosomes takes place during the two mitoses 
by which the mother-cell gives rise to four megaspores. The figures 
are much larger than in the corresponding mitoses in spermatogenesis 
but so much more tedious to secure that most studies in reduction 
have been based upon divisions in the pollen mother-cell. Lilium is 
quite favorable for a study of oogenesis, but it must be remembered 
that it is exceptional in having an embryo sac formed from four 
megaspores. 

In very young stages, before the appearance of the integument, 
the ovary may be removed from the flower and placed directly in the 


304 


METHODS IN PLANT HISTOLOGY 


fixing agent, but in later stages, such as are shown in Figure 113, strips 
should be cut off from the sides of the ovary in order to secure more 
rapid fixing and more perfect infiltration with paraffin. The dotted 
lines in Figure 111C, show about how much should be cut off. This 
is a much better plan than to secure rapid fixing and infiltration by 
cutting the ovary into short pieces, because the ovules will be in 



Fig. 108 .—Lilium canadense: photomicrograph of part of transverse section of ovary, 
showing longitudinal section of ovule with megaspore mother-cell, just before the formation of 
integuments. Fixed in chromo-acetic acid and stained in iron-alum haematoxylin. Eastman 
Commercial Ortho film, Wratten H filter (blue); Spencer 4-mm. objective, N.A. .66; ocular X6; 
arc light; exposure, 2 seconds. Negative by Dr. P. J. Sedgwick. X400. 

about the same stage cf development throughout the ovary, and when 
one finds desirable stages like those from which these photomicro¬ 
graphs were taken, it is gratifying to have these pieces as long as 
possible. 

Chromo-acetic acid, with the addition of a little osmic acid, 
is good for fixing the entire series. Iron-haematoxylin, with a 
light touch of orange, is best for the chromatin. For general beauty 




SPERMATOPHYTES—ANGIOSPERMS 


305 


and for the achromatic structures, the safranin, gentian-violet, 
orange combination has not been excelled. The photomicrographs 
(Figs. 108-110) illustrating the series from the archesporial cell 
(which, in this case, is also the primary sporogenous cell and the 
megaspore mother-cell) to the four megaspore nuclei will repay a 
careful study. One more mitosis produces the 8-nucleate embryo sac, 
but Lilium is not a good type for illustrative purposes, since the egg 
apparatus is not very definitely organized. 



Fig. 109 .—Lilium philadelphicum: photomicrograph of transverse section of ovary showing, 
in one of the ovules on the left, the first mitosis in the megaspore mother-cell; and, in one of the 
ovules on the right, the second mitosis which gives rise to the four megaspore nuclei—chromo- 
acetic acid; safranin, gentian-violet, orange. Cramer contrast plate; 16 mm.-objective; ocular 
X4; yellowish-green filter and also a strong filter such as is used in outdoor work; camera bellows, 
30 cm.; exposure, 2 minutes. Negative by Miss Ethel Thomas. X64. 

For the embryo sac at the fertilization stage, many of the 
Compositae are good. Senecio aureus is quite favorable, because it is 
easy to cut and the akenes do not spread. Aster gives an exceptional 
view of the antipodal region, but is rather hard to cut. Before 
fixing, trim the head as indicated in Figure 111. Silphium, espe¬ 
cially S. laciniatum, furnishes an ideal view of the embryo sac. 
With thumbs and fingers grasp the two wings of the akene and care¬ 
fully split it, exposing the single white ovule inside. This is rather 








306 


METHODS IN PLANT HISTOLOGY 


tedious, but every ovule will yield a perfectly median longitudinal 
section of the embryo sac, and there is not the slightest difficulty in 
cutting. When the rays look their best, the embryo sac is ready for 
fertilization, or the pollen tubes may be entering; as the rays begin 
to wither, you will find fertilization or early stages in the embryo and 
endosperm. Sections should be about 10 n thick. 



Fig. 110 .—Lilium philadelphicum: photomicrograph of second mitosis in megaspore mother¬ 
cell—chromo-aceticacid; safranin, gentian-violet, orange. Cramer contrast plate; 4-mm. objective; 
ocular X4; Abb6 condenser camera bellows, 1 meter; yellowish-green filter and also a strong 
filter such as is used in outdoor work; camera bellows, 1 meter; exposure, 7 minutes. Negative 
by Miss Ethel Thomas. X626. 

The Ranunculaceae, especially Anemone patens var. Wolfgang - 
tana , show a rather large, broad embryo sac, with highly organized 
egg apparatus and antipodals. Sections should be 10 to 20 thick. 

For general views of the embryo sac, the safranin, gentian-violet, 
orange combination is recommended. 




SPERMATOPHYTES—ANGIOSPERMS 


307 


The peculiar embryo sac of Plumbagella, with only two nuclei, 
the egg nucleus and the endosperm nucleus, when ready for fertili¬ 
zation, is shown in Figure 112. 

Fertilization.—’The later stages cut to show the mature embryo 
sac will often show fertilization. The male and female nuclei almost 
invariably show a difference in staining capacity when the male 
nuclei are just discharged 
f rom the pollen tube. With 
cyanin and erythrosin, the 
male nucleus stains blue and 
the female red; hence the 
obsolete terms cyanophilous 
and erythrophilous. As the 
nuclei come into contact 
within the egg, they begin 
to stain alike, the male 
nucleus staining more and 
more like the female. In 
the final stages of fusion it 
is difficult, or impossible, to 
distinguish the two nuclei. 

The male nucleus which 
takes part in the “triple 
fusion” to form the endo¬ 
sperm nucleus behaves in 
the same way. 

Lilium is a very good 
and always available type 
for illustrating fertilization 
(Fig. 113). Take ovaries 
from flowers whose petals have withered but have not yet fallen off. 
Though their embryo sacs and nuclei are smaller, Silphium and Heli- 
anthus are good types, because their curved or twisted male nuclei 
are easily distinguished from the spherical nuclei in the embryo sac. 
The embryo sacs of orchids are very small, but ovules are extremely 
numerous and the chances for securing the fusion of nuclei are cor¬ 
respondingly good. In Cypripedium the nuclei do not fuse in the 
resting condition, but the chromosomes of the two parents are per¬ 
fectly distinct in the egg. The general statement that nuclei fuse 



Fig. 111.— A, head of Aster; B, pod of Capsella; 
C, transverse section of ovary of Lilium. The dotted 
lines show how the material should be trimmed before 
fixing. 





308 


METHODS IN PLANT HISTOLOGY 


in the “ resting condition” is not correct, and probably the chromo¬ 
somes of the two gametes never fuse. 

The Endosperm.—Some of the preparations intended for fertili¬ 
zation will be likely to show early stages in the development of 
endosperm. 



Fig. 112 .—Plumbagella micrantha: longitudinal section of ovule showing the embryo sac, 
with the egg and endosperm nucleus ready for fertilization. Stained in iron-alum haematoxylin. 
Eastman Commercial Ortho film, Wratten E filter (orange); Bausch and Lomb 8-mm. objective 
N.A. .50; Spencer ocular X6; arc light; exposure, 1 second. Preparation by Dr. K. von O. Dahl- 
gren and negative by Dr. P. J. Sedgwick. X208. 

In rather long, narrow embryo sacs, a cell wall is likely to follow 
even the first division of the endosperm nucleus, so that the endo¬ 
sperm is cellular from the beginning. Ceratophyllum, Monotropa, 
and Verbena will furnish material of this type. 

In large, broad embryo sacs, the formation of endosperm is 
almost sure to be initiated by a series of simultaneous free nuclear 




SPERMATOPHYTES—ANGIOSPERMS 


309 



divisions. In large sacs walls then begin to appear at the periphery 
and wall formation gradually advances toward the center until the 
entire sac is filled with tissue. Lilium, Peperomia, and Ranunnlus 
furnish examples of this type. 

An intermediate condition is seen in somewhat elongated embryo 
sacs of medium size, like those of Compositae. After a few free 
nuclear divisions, walls ap¬ 
pear simultaneously through¬ 
out the entire sac. Silphium 
laciniatum is particularly 
good. Akenes from which 
the corolla has just fallen will 
furnish material. 

The Embryo. —In most 
angiosperms the endosperm 
divides earlier than the fertil¬ 
ized egg and in some cases, 
like Asclepias and Casuarina , 
the free nuclear stage of the 
endosperm is completed and 
the cellular stage is well 
advanced before the first di¬ 
vision of the egg. In some 
forms, like the aroids, the 
embryo is massive and differ¬ 
entiation into dermatogen, 
periblem, and plerome comes 
comparatively late; while in 
others, like the Cruciferae, the 
differentiation occurs very 
early. Capsella is a standard example of the latter type (Fig. 114). 
The stages shown in Figure 114 A-F t will be found in pods about 3 mm. 
in length. These may be put directly into the fixing agent, but stages 
like G and H , which are found in pods about 5 mm. in length, should 
be trimmed as indicated in Figure 111#, before fixing. Formalin- 
alcohol-acetic acid is a good fixing agent; the special chromo-acetic- 
osmic acid, with 2 c.c. of acetic acid instead of 3 c.c., is also very 
good, and Delafield’s haematoxylin stains better after the chromic 
series. Cut 5 to 10 p thick and parallel to the flat face of the pod. 


Fig. 113 .—Lilium philadelphicum: photomicro¬ 
graph of section showing fertilization and also the 
triple fusion; from a preparation and negative by 
Dr. W. J. G. Land. X585. 






310 


METHODS IN PLANT HISTOLOGY 


For a study of the monocotyl embryo, Iris, and especially 7. pseuda- 
corus, can be recommended. The embryo is straight, and cotyledon, 
stem-tip, and root are clearly differentiated before the endosperm 



Fig. 114.— Capsella bursa pastoris: A, filamentous stage; B, first division of the embryo cell; 
C, octant stage; D, dermatogen has been cut off, except at the tip of the root; E, differentiation 
into periblem and plerome of the root and differentiation of the periblem of the root; F, further 
development of root but stem not yet differentiated into periblem and plerome; G, completion of 
dermatogen of root; H, two layers of root cap have been formed; H\ topography of entire embryo 
at stage shown in H. Only dermatogen and plerome shaded. A-H, X580; H' X75. 

becomes too hard to cut in paraffin. Fix pieces about 3 mm. wide 
cut perpendicular to the face of the cheese-shaped seed. Do not try 
to cut the whole pod. 





















































SPERMATOPHYTES—ANGIOSPERMS 


311 


Sagittaria has been used quite extensively. It is easily obtained, 
the whole head can be cut with ease, even after the cotyledon and 
stem-tip are clearly differentiated, and the endosperm is instructive; 
but the embryo is curved, like that of Capsella, and good views are 
rather rare. 

Zea Mays , especially the sweet corn, is a good type to illustrate 
the peculiar embryo of the grasses. Directions have been given on 
page 294. 

In many forms good preparations of late stages may be secured 
by soaking the seeds in water until the embryo bursts the seed coat. 
Young seedlings furnish valuable material for a study of vascular 
anatomy. 

Parthenogenesis. —Many embryos are developed without fertili¬ 
zation. Taraxacum , the common dandelion, is an omnipresent 
example. Other widely distributed illustrations are Hieracium 
and practically all species of the Eualchemilla section of the genus 
Alchemilla. Parthenogenetic forms show various irregularities in the 
mitoses leading up to the formation of the egg. 


CHAPTER XXVI 

USING THE MICROSCOPE 

The investigator who desires to see all that his microscope is 
capable of showing must study the optics of his instrument. The 
fundamental principles are presented in any good textbook of physics. 
Excellent practical hints are given in two booklets published by the 
leading American optical companies. These booklets tell the begin¬ 
ner how to set up the microscope, how to keep it in order, and give 
directions concerning illumination, dry and immersion objectives, 
mirror, condenser, diaphragm, and various other things (Fig. 115). 
They were doubtless written for advertising purposes, but since they 
advertise by giving directions for securing the best results with the 
microscope, the information is very reliable. The Spencer Lens Com¬ 
pany of Buffalo, New York, and the Bausch & Lomb Optical Company, 
of Rochester, New York, furnish these booklets free of charge. 

In the histological laboratory where preparations are being made 
the microscope is in constant danger. A cheap microscope with a 
16-mm. objective and one ocular, such an instrument as can be got 
new for $30 or less, can be used for examining preparations while they 
are wet with alcohols, oils, or other reagents. If it is necessary to use 
a better instrument for such work, cover the stage with a piece of 
glass—an old lantern slide is of about the right size—and be extremely 
careful not to get reagents upon the brass portions. 

MICROMETRY 

Everyone who expects to become at all proficient in the use of 
the microscope should learn to measure microscopic objects and 
should learn to form some estimate even without measuring, just 
as one guesses at the size of larger objects. In any measurement 
one should note the tube length, which is usually 160 mm. Since 
the use of the nosepiece is universal, it is convenient to have the 
length measure 160 mm. when the tube is pushed in. Some com¬ 
panies still make the tube so short that it must be pulled out about 
15 mm. to reach the length of 160 mm., even when the nosepiece is 
in place. Where there is no revolving nosepiece, the draw-tube 

312 


USING THE MICROSCOPE 


313 


is simply pulled out until the length is 160 mm. (Fig. 116). Where 
a nosepiece is used, its height should be measured, and the draw-tube 



Body Tube — _ 


Nosepiece .—f - 




Objectives:- 


OcULRR. 


Drbw-Tube 


Graduated ^>hort 
Slide.. 
"Revolvi 
Strg,e-t 

Adjustable^.,,, 

SPRING FlNC,ER. 
Condense* Mou 

Lowe* IrisTi 
for Oblique Lic^ht 

5tace Centering, 
„ Sotiws. 
IRROR,- 

Mirror Forn- 

Mirror Bar..- 

P A c *- & Tin ion 
Button for 
Substf\ce. 


-$ Pinion 

Course Adjustment. 


Fine A^ JUSTMENT 
Buttons 


-Concentric 
Rd justin ^Buttons. 

Lonq 

Slide- 
-Arm 


— Inclination 

Joint 


Pillar. 


— Horseshoe 
3ase. 


Fig. 115.—A modern microscope 

should be pushed in a distance equal to the length of the nosepiece. 
There are in general use two practical methods of measuring micro¬ 
scopic objects, one by means of the ocular micrometer, and the other 
by means of camera lucida sketches. 






314 


METHODS IN PLANT HISTOLOGY 


Measuring with the Ocular Micrometer.—A stage micrometer 
and an ocular micrometer are necessary. A stage micrometer should 
be ruled in tenths and one-hundredths of a millimeter. It does not 
matter what the spacing in the ocular micrometer may be, except 

that the lines must be at equal 
distances from one another. 
As a matter of fact, the ocular 
micrometer is generally ruled 
in tenths of a millimeter, but 
this ruling is more'or less mag¬ 
nified by the lens of the ocular. 

Place the stage micrometer 
upon the stage and the ocular 
micrometer in the tube, and 
arrange the two sets of rulings 
so that ‘the first line in the 
ocular micrometer will coincide 
with the first line of the stage 
micrometer, and then find the 
value of one space in the ocular 
micrometer. The method of 
finding this value is shown in 
the following case in which the 
tube length was 160 mm., the 
ocular a Zeiss ocular micro¬ 
meter 2, and the objective a 
Leitz 3. In the ocular micro¬ 
meter, ninety-eight spaces 
covered just fifteen of the larger 
spaces of the stage micrometer. 
Since the stage micrometer 
is ruled in tenths and one- 
hundredths of a millimeter, the 
fifteen spaces equal 1.5 mm., or 
1,500 /i. 1 Then ninety-eight spaces of the ocular micrometer equal 
1,500 ju; and one space in the ocular equals -fa of 1,500 /jl, or 15.3 /*• 
This value being determined, there is no further use for the stage 
micrometer. To measure the diameter of a pollen grain put the 

1 One millimeter = 1,000 p. The Greek letter p is an abbreviation for pinp6v, or micron. 



































USING THE MICROSCOPE * 


315 


preparation on the stage, using the same objective and ocular micro¬ 
meter, and note how many spaces a pollen grain covers. If the pollen 
grain covers five spaces, its diameter is five times 15.3 or 76.5 /i. 
In the same way, the value of a space in the ocular when used with 
the other objectives should be determined. The values for three 


/OVp. iXft PoL*. vi of 


50 \on 


&hcf< 7 ^Ob- SroL 


Srcb/^ ‘jcc^ 


A 

* 5*0 


A 

a ^ 


Spencer 16mm. 0c.x6. 

Diameter of Field, I700ti.,l.7mm. 

XI02 




Drawn at level of fable 
MirrorHarat HO 


II 

xt § 




Mirror at 05° 

■ill* 

> ^ .1 

. 'NfS 


09/1 X 

■foot mj)° j n jwv //r 

OlX?Q oidojeoua/^ 'umi? zjuon/jjou/jdp 


■dOh 


■*0£ 


riot 


riot ^S 
\b»Lt \httl 
111! 1 ) 11 1 


/cc\cfi 


§ ^ 
I 

Co X. 


-c „ 
?- 


r 

r 

-r 

?- 


* T 
2 


Fig. 117.—Card scale for practical use 


or four objectives may be written upon an ordinary slide label and 
pasted upon the base of the microscope for convenient reference. 

This method is the best one for measuring spores and for most 
measurements in taxonomy. 

Measuring by Means of Camera Lucida Sketches. —This method 
is of great importance in research work, because various details can 







316 


METHODS IN PLANT HISTOLOGY 


be measured with far greater rapidity than by the other method. 
Upon a piece of cardboard, about as thick as a postal card, draw a 
series of scales like those shown in Figure 117. 

Make a scale for each objective. It is not necessary to make 
scales for all the oculars, but only for the one in most constant use. 
It is absolutely necessary to note the tube length, length of the bar 
of the camera mirror and inclination of the camera mirror, and the 
level at which the scale is made. A variation in any of these details 
will change the scale. 

In using the stage micrometer, place the cardboard on the table, 
and with the aid of the camera lucida sketch the rulings of the microm¬ 
eter. In Figure 117 note, for example, the scale drawn with Spencer 
16 mm. objective, ocular X6. The spaces are drawn from the tenths 
of a millimeter rulings of the stage micrometer. Therefore each space 
on the card represents one-tenth of a millimeter or 100 n, and the 
ten spaces shown on the card represent 1 mm., or 1,000 /*. By measur¬ 
ing with a metric rule the ten spaces upon the card, it is found that the 
scale is 102 mm. in length. The magnification of any drawing made 
with the same ocular and objective, under the same conditions, will 
therefore be 102 diameters. This does not mean that the magnifying 
power is 102 diameters, for the magnification of this combination is 
much less. A scale drawn at the level of the stage would show more 
nearly the magnifying power of the combination, but would still give 
too large a figure. The exact size of any object which has been 
sketched with this combination can now be measured by applying 
the cardboard scale, just as one would measure gross objects with a 
rule. 

The diameter of the field with this-combination is 1,700 ju* By 
knowing the diameter of the field with the various combinations, one 
can guess approximately the size of objects. 

Other combinations are made in the same way. An excellent 
check on the accuracy of the computations is to measure the same 
object by means of the ocular micrometer and by the scale card. If 
the results are the same, the computations are correct. 

In making sketches, it is a good plan to add the data which would 
be needed at any time in making measurements; e.g., Spencer objec¬ 
tive 16 mm., ocular X6, table, 110, 45°, would show that the sketch 
was made at the level of the table, with the mirror bar at 110, and 
the camera mirror at an angle of 45.° 


USING THE MICROSCOPE 


317 


ARTIFICIAL LIGHT 

During a considerable part of the year daylight is often insufficient 
for successful work with the microscope. Numerous contrivances 
for artificial illumination have been devised, some of them fairly 
good, but most of them thoroughly unsatisfactory. More than two 
hundred years ago Hooke used a device for artificial illumination 
which probably suggested the apparatus used by the late Professor 
Strasburger at Bonn. The apparatus in use in our own laboratory is 
only a slightly modified form of that used in the Bonn laboratory. 

The apparatus consists, essentially, of a hollow sphere filled with 
liquid. A fairly good and practical light can be got with an ordinary 
lamp by allowing the light to pass through a wash bottle filled with a 
weak solution of ammonia copper sulphate. A piece of dark paper 
with a circular hole in it serves as a diaphragm, and at the same time 
protects the eyes from the direct light of the lamp. 

At present, we.are using a white 50-watt, 115-volt, nitrogen Mazda 
bulb, with a shade to protect the eyes. This not only furnishes a 
strong light, without any glare, but throws a good light on the pencil, 
which is an important consideration in drawing with a camera lucida. 

Optical companies are now making excellent lights for microscopes. 
These lights furnish good illumination and most of them have the 
effect of good daylight. 

If laboratory tables are small, seating only one student, there 
should be a plug to attach the table to some convenient outlet; and 
also another outlet on the table for the microscope lamp. If the 
table is large, seating four or more students, there should be an 
outlet on the table for each student, and a single plug by which the 
whole table may be connected with a convenient outlet. 

For elementary classes, which are not likely to use higher powers 
than a 4 mm. objective with an ocular magnifying five or six times, 
individual lamps are not necessary in a well-lighted laboratory. 
Half-a-dozen strong lights, of the overhead type, with white shades, 
serve very well for a class of twenty-five or thirty students. 


CHAPTER XXVII 

LABELING AND CATALOGUING PREPARATIONS 
THE LABEL 

We should say that the first thing to write upon a label is the 
genus and species of the plant; the next thing would be the name of 
the organ or tissue, and then might be added the date of collection, 
e.g., Marchanita polymorpha , young archegonia, January 10, 1915. 
The date of making the preparation is of no value unless the student 
is testing the permanence of stains or something of that sort. It 
is hardly worth while to write upon the label the names of the stains 
used, for the student will soon learn to recognize the principal stains. 
A hasty sketch on the label will often indicate any exceptionally 
interesting feature in the preparation. To facilitate finding such a 
feature, it is a good plan to mark the particular section or sections with 
ink, the marking being always on the underside of the slide so as 
not to cause any inconvenience if an immersion lens should be used. 

CATALOGUING PREPARATIONS 

As a collection grows, the student will need some device for 
locating readily any particular preparation. Some have their slides 
numbered and catalogued, but all devices of this sort are too cumbrous 
and slow for the practical worker in the laboratory. After thirty 
years’ experience with a collection which now numbers more than 
thirty thousand preparations, we recommend the following system: 

Four wooden slide boxes of the usual type will do for a beginning; 
they should be labeled: Thallophytes, Bryophytes, Pterido- 
phytes, and Spermatophytes. As the collection grows and new 
boxes are needed, the classification can be made more definite, e.g., 
there should be a box labeled Bryophytes Hepaticae and one labeled 
Bryophytes Musci. As the liverwort collection grows, three boxes 
will be necessary, and should be labeled Bryophytes Hepaticae 
Marchantiales, Bryophytes Hepaticae Jungermanniales, and Bryo¬ 
phytes Hepaticae Anthocerotales. It will readily be seen that the 
process can be continued almost indefinitely, and that new slides 

318 


LABELING AND CATALOGUING PREPARATIONS 319 

may be at any time dropped into their proper places. A rather com¬ 
plete label gradually built up in this way is shown in Figure 118. 

The beginner will often find that the mere placing of a slide in the 
proper box and the box in its proper place on the shelf will refresh or 


BRYOPHYTES 

HEPATICAE 

Jungermann idles 

Porella platyphyllum 

Archegonia 

Fig. 118.—Label for slide box 

increase his knowledge of classification. While this system is almost 
ideal for the careful worker, especially if he has some knowledge of 
classification, it is the worst possible method for a careless student 
since a slide in the wrong box is almost hopelessly lost if the collec¬ 
tion is large enough to need thorough classifying. 



CHAPTER XXVIII 

A CLASS LIST OF PREPARATIONS 

Where a regular course in histology is conducted, it is a good plan 
to give each student at the outset a complete list of the preparations 
which he is expected to make. In a three months’ course a fairly 
representative collection of preparations can be made. The avail¬ 
ability of material determines what a list shall be. Besides gaining 
an introduction to the use of the microscope and its accessories, a class 
meeting ten hours a week for twelve weeks should be able to do as 
much work as is outlined below. 

In making the mounts, the order indicated in the list should not 
be followed. Begin with temporary mounts, and then study, in 
succession, freehand sections (the glycerin method), the Venetian 
turpentine method, the paraffin method (the celloidin method), and 
special methods. A large proportion of the time should be devoted 
to the paraffin method. 

It is neither possible nor desirable that each student should in 
every case go through all the processes from collecting material to 
labeling. Some of the material may be in 85 per cent alcohol, some 
in formalin, some in glycerin, some in Venetian turpentine, and some 
in paraffin. One student may imbed in paraffin enough of the 
Anemone for the whole class; another may imbed the Lilium stamens; 
and by such a division of labor a great variety of preparations may 
be secured without a corresponding demand upon the time of the 
individual. 

LIST OF PREPARATIONS 

THALLOPHYTES 

SCHIZOPHYTES 

MYXOMYCETES 

1. Trichia varia .—Paraffin sections, 5 (jl. Safranin, gentian-violet, orange. 

SCHIZOMYCETES 

2. Bacteria. — Coccus, Bacillus, and Spirillum forms. Stain on cover-glass 

or slide. 


320 


A CLASS LIST OF PREPARATIONS 


321 


3. Bacillus anthracis. —In liver of mouse. Paraffin sections, 5 p. Stain 
in gentian-violet, Gram’s method. 

4. Oscillatoria. —Fix in special chromo-acetic-osmic acid and stain in iron- 
alum haematoxylin to show nuclei. Venetian turpentine method. 

5. Tolypothrix. —Use the Venetian turpentine method. Should show 
heterocysts, hormogonia, and false branching. 

6. Nostoc. —Venetian turpentine method. 

7. Wasserblicthe: —The principal forms in this material are: 

a) Coelosphaerium Kutzingianum. —Colonies in the form of hollow 
spheres. 

b) Anabaena gigantea. —Filaments straight. Preparations should show 
vegetative cells, heterocysts, hormogonia, and spores. 

c) Anabaena flos-aquae .—Filaments curved. Stain on the slide and 
mount in balsam. If material is abundant, stain in iron-alum 
haematoxylin and mount in Venetian turpentine. 

8. Gloeotrichia. —Smear on the slide, stain in safranin and gentian-violet, 
and mount in balsam; or use the Venetian turpentine method, staining 
in Magdala red and anilin blue and crushing under the cover-glass. 

ALGAE 

CHLOROPHYCEAE 

9. Volvox. —Use the Venetian turpentine method. If paraffin material 
is available, cut 5p in thickness and stain in safranin, gentian-violet, 
orange. 

10. Scenedesmus. —Let a drop containing the material dry upon the slide 
stain, and mount in balsam. 

11. Hydrodictyon. —Use the Venetian turpentine method. 

Each preparation should contain pieces of old and of young nets, and 
also at least one young net developing within an older segment. The 
greatest care must be taken not to injure the older segments while arran¬ 
ging the mount. 

12. Ulothrix. —Use the Venetian turpentine method. Each mount should 
show various stages in the development of spores and gametes. 

13. Oedogonium .—Stain in Magdala red and anilin blue and mount in 
Venetian turpentine. 

14. Coleochaete. —Stain in Delafield’s haematoxylin and mount in balsam. 

15. Cladophora. —Stain some in iron-haematoxylin and some in Magdala 
red and anilin blue. Mount both together in Venetian turpentine. 

16. Diatoms. —Make mounts of the frustules and also stained preparations 
showing the cell contents. 

17. Desmids. —Make mounts of available forms. Use the Venetian turpen¬ 
tine method if material is sufficiently abundant. 


322 METHODS IN PLANT HISTOLOGY 

18. Zygnema. —Stain in iron-haematoxylin, and mount in Venetian turpen¬ 
tine. 

19. Spirogyra. —Stain in Magdala red and anilin blue, and mount in Venetian 
turpentine. 

20. Vaucheria. —Stain in iron-alum haematoxylin, and mount in Venetian 
turpentine. 

21. Char a. —Cut paraffin sections of the apical cell, oogonia, and antheridia. 

PHAEOPHYCEAE 

22. Ectocarpus — Stain some in iron-haematoxylin and some in Magdala 
red and anilin blue. Mount both together in Venetian turpentine. 

23. Cutleria. —Sections of oogonia, antheridia, and sporangia. Cut 10 /a 
thick and stain in iron-haematoxylin with about 7 minutes in safranin. 

24. Fucus vesiculosus . —Antheridial conceptacle with paraphyses and 
antheridia; oogonial conceptacle with oogonia. Cut 10 p thick and 
stain in iron-haematoxylin with about 5 minutes in safranin. 

RHODOPHYCEAE 

25. Nemalion. —Stain some in iron-haematoxylin and some in eosin. Each 
preparation should show trichogyne, carpogonium, and cystocarp. You 
cannot mount it in Venetian turpentine; use glycerin or glycerin jelly. 

26. Polysiphonia. —Stain in iron-haematoxylin or Magdala red and anilin 
blue. Mount whole in Venetian turpentine. Each mount should 
show tetraspores, antheridia, and cystocarps. If material is in paraffin, 
cut sections about 7 p thick. 

FUNGI 

PHYCOMYCETES 

27. Rhizopus nigricans. —Stain young sporangia in eosin, dilute Delafield’s 
haematoxylin, or in Magdala red and anilin blue. Some zygosporic 
material should be stained strongly in eosin: some should be stained 
in iron-alum haematoxylin, and in reducing the stain, some should 
be taken out early to show the coenocytic character of the mycelium, 
and some should be drawn farther to show the structure of the zygospores 
and suspensors. Venetian turpentine. 

28. Saprolegnia. —Stain some in Magdala red and anilin blue, and some in 
iron-alum haematoxylin. Each mount should show sporangia and 
oogonia. Venetian turpentine. 

29. Albugo Candida. —Select white blisters which have not yet broken open. 
Paraffin 8 p. Iron-alum haematoxylin and orange. Oogonia and 
antheridia, 5 p, same stain. 

30. Albugo bliti on Amarantus retroflexus. —Cut out small portions of leaves 
in which the oogonia can be seen in abundance. Paraffin, 5 p. 


A CLASS LIST OF PREPARATIONS 


323 


ASCOMYCETES 

31. Peziza. —Paraffin sections of young apothecia, 5 p or less; sections of 
older apothecia, 10 or 15 /x. Safranin, gentian-violet, orange. 

32. Aspergillus, Eurotium. —Stain in eosin and mount in Venetian turpentine. 

33. Penidllium. —Treat like Aspergillus. 

34. Erysiphe commune on Polygonum aviculare. —Strip the fungus from the 
leaf. Paraffin, 5 p or less. Safranin, gentian-violet, orange. 

35. Uncinula necator on Ampelopsis quinquefolia. —Stain in Magdala red 
and anilin blue. Mount whole in Venetian turpentine and break the 
perithecia under the cover. 

36. Xylaria. —Paraffin sections of younger stages. Delafield’s haematoxylin 
and erythrosin. Be sure that some section in each mount shows the 
opening of a perithecium. 

LICHENS 

37. Physcia stellaris. —Cut in paraffin, 5 p. Stain in cyanin and erythrosin. 

BASIDIOMYCETES 

38. Puccinia graminis. —Aecidium stage on barberry leaf. Uredospore 
and teleutospore stage on oats. Cut 3 p and stain in iron-haematoxylin. 

39. Coprinus micaceus. —Paraffin. Transverse sections of gills showing 
trama, paraphyses, basidia, and spores. To show the basidium with 
four spores, the sections should be 15 p thick. For development of the 
spores, cut 5 p or less. Safranin, gentian-violet, orange. Boletus, 
Hydnum, and Polyporus are treated in the same manner. 

BRYOPHYTES 

HEPATICAE 

40. Riccia natans. —Paraffin, 10 or 15 p. Delafield's haematoxylin. Arche- 
gonia, antheridia, and sporophytes imbedded in the gametophyte. 

41. Marchantia polymorpha. —Paraffin, 5 or 10 p. Archegonia, antheridia, 
and sporophytes. 

42. Anthoceros laevis. —Paraffin, 5 or 10 p. Longitudinal and transverse 
. sections of sporophyte. Safranin, gentian-violet, orange. 

43. Pellia epiphylla. —Paraffin, 5 or 10 p. Longitudinal sections of sporo¬ 
phyte attached to gametophyte. Safranin, gentian-violet, orange. 

44. Porella platyphylla.—Psirafim, 10 p. Delafield’s haematoxylin. Arche¬ 
gonia, antheridia, sporophyte, and apical cell. 

MUSCI 

45. Sphagnum. —Leaf buds. Cut 5 p and stain in safranin and anilin blue. 

46. Sphagnum. —Capsule. Paraffin. Delafield’s haematoxylin and ery¬ 
throsin. 


324 


METHODS IN PLANT HISTOLOGY 


47. Funaria hygrometrica. —Paraffin. Longitudinal and transverse sections 
of young capsules. Delafield’s haematoxylin. 

48. Funaria hygrometrica or any favorable form. Protonema. Place the 
well-cleaned material directly into 50 per cent glycerin and allow it to 
concentrate. Mount in glycerin or glycerin jelly. 

49. Bryum. —Paraffin. Antheridia, 10 p; archegonia, 15 to 20 p; capsule, 
lO/i. 


PTERIDOPHYTES 

LYCOPODIALES 

50. Lycopodium lucidulum. —Transverse section of stem. Paraffin sections. 
Safranin and Delafield’s haematoxylin. 

51. Lycopodium inundatum. —Paraffin. Longitudinal sections of strobilus. 

52. Selaginella. —Paraffin. Longitudinal sections of rather mature strobili. 
Safranin, gentian-violet, orange. 

53. Isoetes echinospora. —Transverse section of stem. Paraffin. Safranin 
and Delafield’s haematoxylin. 

54. Isoetes echinospora. —Paraffin. Longitudinal sections of microsporangia 
and megasporangia. Safranin, gentian-violet, orange. 

EQUISETALES 

55. Equisetum arvense. —Prothallia in Venetian turpentine. Stem-tips in 
paraffin. Transverse section of stem freehand or in celloidin. 

FILICALES 

56. Botrychium virginianum. —Paraffin. Stain rhizome, stipes, and root 
in safranin and Delafield’s haematoxylin. Stain sporangia in iron- 
haematoxylin. 

57. Protostele. —Use Gleichenia. Cut freehand and stain in safranin and 
anilin blue. 

58. Solenostele (amphiphloic siphonostele). —Use Adiantum or Dicksonia 
punctilobula. 

59. Ectophloic siphonostele. —Use Osmunda cinnamomea. 

60. Polystele. —Use Pteris aquilina or any species of Polypodium. 

61. Sporangia. —For development, use Pteris, Aspidium, Cyrtomium, or 
try any available species. For mitosis, Osmunda is exceptionally good. 

62. Antheridia and archegonia. —Mount whole in Venetian turpentine. 
Iron-alum haematoxylin. Sections should be 5 to 10 p thick. Stain in 
iron-haematoxylin and orange. 

63. Embryo—Pteris and Adiantum are good. Cut longitudinal vertical 
sections 10 p thick. 


A CLASS LIST OF PREPARATIONS 


325 


SPERMATOPHYTES 

GYM NOSPERMS 

CYCADALES 

64. Zamia. —Freehand sections of stem. Safranin and light green. Trans¬ 
verse sections of microsporophyll, 5 or 10/z. Longitudinal sections of 
entire ovule, 10 to 15 ^u; stain in safranin, gentian-violet, orange. 
Longitudinal sections of nucellus with pollen tubes, 10 /z. Iron-haema- 
toxylin and orange. 

GINKGOALES 

65. Ginkgo biloba. —Longitudinal sections of endosperm showing archegonia 
or young embryos. Paraffin 10 p. 

Sections of microsporangia with nearly mature pollen, 5 /z. 

CONIFERALES 

66. Pinus Laricio. —Transverse sections of needles and young stem. Free¬ 
hand. Safranin and light green. 

67. Pinus Strobus. —Freehand sections of well-seasoned wood. Safranin 
and Delafield’s haematoxylin. 

68. Pinus Laricio. —Paraffin. Longitudinal section of mature staminate 
strobilus. Safranin, gentian-violet, orange. 

69. Abies balsamea or Pinus Laricio. —Pollen at shedding stage shaken out 
and imbedded in paraffin; 5 p. Stain in safranin, gentian-violet, orange. 

70. Pinus Laricio. —Paraffin. Ovule with archegonia. Safranin, gentian- 
violet, orange. 

71. Pinus sylvestris or P. Laricio. —Paraffin. Embryos. Cyanin and ery- 
throsin, or safranin and anilin blue. 

GNETALES 

72. Transverse section of stem of Ephedra. Freehand. 

73. Longitudinal section of the ovule of Ephedra. 

A NGIO SPERM S 
DICOTYLS 

74. Pelargonium. —Transverse sections of stem to show phellogen and 
intrafascicular cambium. Freehand. Endarch siphonostele. 

75. Tilia americana. —Celloidin or freehand. Transverse sections of small 
stems 3 mm. to 6 mm. in diameter. Safranin and Delafield’s haematoxy¬ 
lin. Endarch siphonostele with annual rings. 

76. Sambucus nigra. —Transverse section of primary root to show origin 
of secondary structures. 

77. Cucurbita. —Longitudinal section of stem to show sieve tubes. 


326 


METHODS IN PLANT HISTOLOGY 


78. Capsella bursa-pastor is. —Paraffin. Floral development, 5 p. Embryos, 
5 to 10 p. Stain both in Delafield’s haematoxylin without any contrast 
stain. 

79. Taraxacum officinale. —Paraffin. Floral development, 5 p. Embryo 
sac, 10 to 15 p. 

80. Ranunculus. —Longitudinal sections of young flowers to show megaspore 
mother-cells and megaspores. 

81. Silphium. —Longitudinal sections of the ovule at the fertilization period. 
Longitudinal sections of staminate flowers just before the shedding of 
pollen. 

82. Anemone patens. —Paraffin. Embryo sac. 

MONOCOTYLS 

83. Clintonia. —Transverse section of stem to show siphonostele. Paraffin. 
Safranin and anilin blue. 

84. Acorus calamus. —Transverse sections of rhizome, freehand or paraffin, 
to show amphivasal bundles. 

85. Zea Mays. —Transverse section of stem to show scattered bundles; also 
good for companion cells. Freehand. Safranin and anilin blue. 

86. Tradescantia virginica. —Longitudinal sections of root-tip. Paraffin, 
5 and 10 p. Stain for mitosis. 

87. Smilax herbacea. —Transverse section of adult root. Freehand. Shows 
exarch, radial structure, and a highly developed endodermis. Safranin 
and Delafield’s haematoxylin. 

88. Lilium. —Transverse section of leaf. Freehand. Transverse section 
of ovaries in various stages from megaspore mother-cell to fertilization; 
transverse sections of anthers to show microspore mother-cells and reduc¬ 
tion of chromosomes; also later stages with nearly mature pollen. 
Paraffin 5 to 10 p. 

89. Iris. —Section of young seeds to show embryo with cotyledon and 
stem-tip. 

90. Sagittaria. —Longitudinal sections of ovulate flowers of various stages 
to show development of the embryo and endosperm. 

91. Zea Mays. —Longitudinal and transverse sections of embryo (sweet corn, 
roasting-ear condition) to show structure of root and beginning of pro- 
toxylem. 


CHAPTER XXIX 


FORMULAS FOR REAGENTS 
FIXING AGENTS 

Absolute Alcohol.— Used alone without any mixtures. 

Carnoy’s Fluid.— 

Absolute alcohol. 2 parts 

Chloroform. 3 parts 

Glacial acetic acid. 1 part 

Farmer’s Fluid.— 

Absolute alcohol. 3 parts 

Glacial acetic acid. 1 part 

Formalin Alcohol (Lynds Jones’s formula). — 

70 per cent alcohol. 100 c.c. 

Commercial formalin. 2c.c. 

Formalin Alcohol (Dr. Land’s formula). — 

50 per cent alcohol. 100 c.c. 

Commercial formalin. 6 c.c. 

Formalin Acetic Alcohol.— 

50 per cent alcohol. 90 c.c. 

Commercial formalin. 5c.c. 

Glacial acetic acid. 5 c.c. 

Formalin Acetic Acid.— 

Commercial formalin. 10 c.c. 

Glacial acetic acid. 5c.c. 

Water. 85 c.c. 

Formalin.— 

Commercial formalin. 3 to 10 c.c. 

Water. 100 c.c. 

Stock Chromo-Acetic Solution.—• 

Chromic acid. 1 g- 

Glacial acetic acid. 1 c.c. 

Water. 100 c.c. 


327 






















328 


METHODS IN PLANT HISTOLOGY 


Schaffner’s Chromo-Acetic Solution.— 

Chromic acid. 0.3 g. 

Glacial acetic acid. 0.7 c.c. 

Water. 99.0 c.c. 

Chromo-Acetic Solution (for marine algae).— 

Chromic acid. 1-0 g. 

Glacial acetic acid. 0.4 c.c. 

Sea-water. 400.0 c.c. 

Material must be washed in sea-water. 

Strong Chromo-Acetic Solution.— 

Chromic acid. 1 g- 

Glacial acetic acid. 3 c.c. 

Water. 100 c.c. 

Licent’s Formula.— 

1 per cent chromic acid. 80 c.c. 

Glacial acetic acid. 5 c.c. 

Formalin. 15 c.c. 

Flemming’s Fluid (weaker solution).— 

f 1 per cent chromic acid (in water). 25 c.c. 

A \ 1 per cent glacial acetic acid (in water). 10 c.c. 

(Water. 55 c.c. 

B. 1 per cent osmic acid (in water).. 10 c.c. 

Keep the mixture A made up, and add B as the reagent is needed 
for use, since it does not keep well. 

Flemming’s Fluid (stronger solution).— 

1 per cent chromic acid. 45 c.c. 

2 per cent osmic acid. 12 c.c. 

Glacial acetic acid. 3 c.c. 

Special Chromo-Acetic-Osmic Solution.— 

Chromic acid. 1 g. 

Glacial acetic acid. 3 c.c. 

1 per cent osmic acid. 1 c.c. 

Water. 100 c.c. 

Good for algae (fresh-water), filamentous fungi, and many 
others; but not for root-tips, stem-tips, and many other things. 
The name, special, was selected because the formula is not general in 
its application. 

























FORMULAS FOR REAGENTS 


329 


Merkel’s Fluid.— 

1.4 per cent solution of chromic acid. 25 c.c. 

1.4 per cent solution of platinic chloride. 25 c.c. 

Benda’s Fluid.—* 

1 per cent chromic acid. 16 c.c. 

2 per cent osmic acid. 4 c.c. 

Glacial acetic acid. 2 drops 

Hermann’s Fluid.— 

1 per cent platinic chloride. 15 parts 

Glacial acetic acid. 1 part 

2 per cent osmic acid. 4 or 2 parts 

Picric Acid.—• 

Picric acid. 1 g. 

Water or 70 per cent alcohol. 100 c.c. 

Bouin’s Fluid.— 

Commercial formalin. 25 c.c. 

Picric acid (saturated solution in water). 75 c.c. 

Glacial acetic acid. 5 c.c. 

Corrosive Sublimate and Acetic Acid.— 

Corrosive sublimate. 3 g. 

Glacial acetic acid. 5 c.c. 

70 per cent alcohol (or water). 100 c.c. 

Corrosive Sublimate, Formalin, Acetic Acid.— 

Corrosive sublimate. 4 g. 

Formalin. 5 c.c. 

Glacial acetic acid. 5 c.c. 

Water (or 50 per cent alcohol). 100 c.c. 

For material to be mounted in Venetian turpentine, use the 
aqueous solution; for imbedding, use the alcoholic. Both this and 
the preceding solution should be used hot (85° C.). 

Bensley’s Formula (for mitochondria).— 

2\ per cent corrosive sublimate in water.4 parts 

2 per cent osmic acid. 1 part 
























330 


METHODS IN PLANT HISTOLOGY 


Corrosive Sublimate, Acetic Acid, and Picric Acid.— 


Corrosive sublimate. 5 g. 

Glacial acetic acid. 5 c.c. 

Picric acid.. • • 1 g- 

50 per cent alcohol.. 100 c.c. 

Corrosive Sublimate and Picric Acid (Jeffrey’s formula).— 
Corrosive sublimate, saturated solution in 30 

per cent alcohol. 3 parts 

Picric acid, saturated in 30 per cent alcohol. 1 part 

Iodine.—• 


To a saturated solution of potassium iodide in distilled water, add iodine 
to saturation. Filter and dilute with distilled water to a rich brown color. 
For fixing, dilute still farther to a light-brown color. Good for unicellular 
and filamentous algae. See directions in chapter ii. 

Gram’s Potassium Iodide Solution (for staining bacteria).— 


Iodine. 1 g- 

Potassium iodide. 2 g. 

Water. 300 c.c. 


Iodine (solution for starch test).—• 

Dissolve 1 -g. potassium iodide in 100 c.c. of water and add 0.3 g. 


iodine. 

Gilson’s Fluid.—• 

95 per cent alcohol. 42 c.c. 

Water. 60 c.c. 

Glacial acetic acid. 18 c.c. 

Concentrated nitric acid. 2 c.c. 

Corrosive sublimate (saturated solution in water) 11 c.c. 

Bensley’s Formula (for canal system).— 

1. Bichromate of potash. g. 

2. Corrosive sublimate.... 5 g. 

3. Water. 90 c.c. 

4. Formalin (neutral). 10 c.c. 

Make a solution of 1 , 2, 3, and then add the neutral formalin. 

Osmic Acid (stock solution).—• 

Osmic acid. 1 c.c. 

Distilled water. lc.c. 


The bottle in which the solution is to be kept, and also the glass 
tube in which the acid is sold, must be thoroughly cleaned. Break 




















FORMULAS FOR REAGENTS 


331 


off the end of the tube, and drop both tube and acid into the distilled 
water, or simply drop the tube into the bottle and shake the bottle 
until the tube breaks. 

Osmic Acid.— 

Five or six drops of the stock solution to 50 c.c. of water is good for 
unicellular and colonial algae. In many cases 1 or 2 c.c. to 100 c.c. of water 
is better. 

STAINS 

Delafield’s Haematoxylin.— 

To 100 c.c. of a saturated solution of ammonia alum add, drop by drop, 
a solution of 1 g. of haematoxylin dissolved in 6 c.c. of absolute alcohol. 
Expose to air and light for one week. Filter. Add 25 c.c. of glycerin and 
25 c.c. of methyl alcohol. Allow to stand until the color is sufficiently dark. 
Filter and keep in a tightly stoppered bottle. 1 

The solution should stand for at least 2 months before it is ready 
for using. 

Erlich’s Haematoxylin.— 


Distilled water. 50 c.c. 

Absolute alcohol. 50 c.c. 

Glycerin. 50 c.c. 

Glacial acetic acid. 5 c.c. 

Haematoxylin. 1 g- 

Alum in excess. 


Keep it in a dark place until the color becomes a deep red. If 
well stoppered, it will keep indefinitely. 

Boehmer’s Haematoxylin.— 


f Haematoxylin. 1 g- 

\ Absolute alcohol. 12 c.c. 

f Alum. 1 g- 

\ Distilled water. 240 c.c. 


The solution A must ripen for 2 months. When wanted for 
use, add about 10 drops of A to 10 c.c. of B. Stain 10 to 20 minutes. 
Wash in water and proceed as usual. 

Mayer’s Haem-Alum. —Haematoxylin, 1 g., dissolved with heat 
in 50 c.c. of 95 per cent alcohol and added to a solution of 50 g. of 


i Stirling and Lee. 











332 


METHODS IN PLANT HISTOLOGY 


alum in a liter of distilled water. Allow the mixture to cool and 
settle; filter; add a crystal of thymol to preserve from mold (Lee). 

It is ready for use as soon as made up. Unless attacked by mold, 
it keeps indefinitely. 

Haidenhain’s Iron-Haematoxylin. —This stain was introduced 
by Haidenhain in 1892 and has gained a well-deserved popularity 
with those engaged in cytological work. Two solutions are used, 
and they are never mixed: 

A. to 4 per cent aqueous solution of ammonia sulphate of iron. Use 
the ferric (violet) crystals, not the ferrous (green) crystals. 

B. f per cent solution of haematoxylin in distilled water. 

The crystals of haematoxylin will dissolve in the distilled water 
in about 10 days; the stain reaches its greatest efficiency in about 
6 weeks. About 3 months from the time it is made up, it begins to 
deteriorate. A stain made by dissolving the crystals in strong alco¬ 
hol and then diluting with water so as to get a practically aqueous 
solution is not so good. 

Greenacher’s Borax Carmine.— 


Carmine. 3 g. 

Borax. 4 g. 

Distilled water. 100 c.c. 


Dissolve the borax in water and add the carmine, which is quickly 
dissolved with the aid of gentle heat. Add 100 c.c. of 70 per cent 
alcohol and filter (Stirling). 

Alum Carmine.— A 4 per cent aqueous solution of ammonia alum 
is boiled 20 minutes with 1 per cent of powdered carmine. Filter 
after it cools (Lee). 


Alum Cochineal.—• 

Powdered cochineal. 50 g. 

Alum. 5 g. 

Distilled water. 500 c.c. 


Dissolve the alum in water, add the cochineal, and boil; evaporate 
down to two-thirds of the original volume and filter. Add a few 
drops of carbolic acid to prevent mold (Stirling). 








FORMULAS FOR REAGENTS 


333 


Piero-Carmine.— 


Picro-carmine (picro-carminate of ammonia). 1 g. 

Water. 100 c.c. 


Myer’s Carmalum.— 


Carminic acid. 1 g. 

Alum. 10 g. 

Distilled water. 200 c.c. 


Dissolve with heat; decant or filter, and add a crystal of thymol 
to avoid mold. 

This is the stain recommended for Volvox. 


Carmalum (Alum Lake).— 


Carmalum. 1 g- 

Water. 100 c.c. 

Ammonia. lc.c. 


Filter, if there is any precipitate. 

Aceto-Carmine.— 

Heat a 45 per cent aqueous solution of glacial acetic acid to the boiling- 
point, with an excess of powdered carmine. Cool and filter. 


Iron Aceto-Carmine.— 

Add a trace of ferric hydrate, dissolve 45 per cent acetic acid, to a 
quantity of acetic carmine until the liquid becomes bluish red, but no precipi¬ 
tate forms. Then add an equal amount of ordinary acetic-carmine. 

Eosin.— 

Eosin. 

Water, or 70 per cent alcohol. 100 c.c. 

General Formula for Anilins. —Make a 3 per cent solution of 
anilin oil in distilled water; shake well and frequently for a day; 
add enough alcohol to make the whole mixture about 20 per cent 
alcohol; add 1 g. of cyanin, erythrosin, safranin, gentian-violet, etc., 
to each 100 c.c. of this solution. 

Cyanin— This general formula is not at all successful with 
Gruber’s cyanin, but gives satisfactory results with an immensely 
cheaper cyanin, sold by H. A. Metz & Company, 122 Hudson Street, 
New York. 











334 


METHODS IN PLANT HISTOLOGY 


Anilin Blue.— 

Anilin blue.•. 1 g. 

85 or 90 per cent alcohol. 100 c.c. 


For staining before mounting in Venetian turpentine, this stain 
should be made up in strong alcohol, even if the dry stain is intended 
for aqueous solution. 


Iodine Green.— 

Iodine green. 1 g. 

70 per cent alcohol. 100 c.c. 

Methyl Green.— 

Methyl green. 1 g. 

Glacial acetic acid. 1 c.c. 

Water. 100 c.c. 


If the preparation is to be mounted in balsam, a slight trace of 
acetic acid and also a trace of methyl green should be added to the 
absolute alcohol used for dehydrating. 

For staining vascular bundles, the acid may be omitted, even from 
the formula. 


Light Green.— 

Light green. 1 g. 

Clove oil. 100 c.c. 

or 

Light green . 1 g. 

Clove oil. 75 c.c. 

Absolute alcohol. 25 c.c. 

Fuchsin.— 

Fuchsin. 1 g. 

95 per cent alcohol. 100 c.c. 

Water. 100 c.c. 

Acid Fuchsin.— 

Acid fuchsin. 1 g. 

Water. 100 c.c. 


Use this formula when staining woody tissues in methyl green 
and acid fuchsin. 



















FORMULAS FOR REAGENTS 


335 


Ziehl’s Carbol Fuchsin.— 

Fuchsin. 1 g. 

Carbolic-acid crystals. 5 g. 

95 per cent alcohol. 10 c.c. 

Water. 100 c.c. 

Fuchsin and Iodine Green Mixtures.—-Two solutions are kept 
separate, since they do not retain their efficiency long after they are 
mixed. 

. (Fuchsin (acid). 0.1 g. 

\ Distilled water. 50.0 c.c. 

-p. (Iodine green...,. 0.1 g. 

\ Distilled water. 50.0 c.c. 

( Absolute alcohol. 100.0 c.c. 

C | Glacial acetic acid. 1.0 c.c. 

[iodine. 0.1 g. 

Stain in equal parts of A and B. Transfer from the stain directly 
to solution C, and from C to xylol. 

Another Formula.— 

. (Acid fuchsin. 0.5 g. 

A \ Water. 100.0 c.c. 

(Iodine green. 0.5 g. 

\ Water. 100.0 c.c. 

Mix a pipette full of A with a pipette full of B; stain 2 to 8 min¬ 

utes; dehydrate rapidly and mount in balsam. 

Magdala Red.— 

Magdala red. 1 g- 

85 or 90 per cent alcohol. 100 c,c. 

Use formula when staining in Magdala red and anilin blue, 
before mounting in Venetian turpentine. Phloxine is the American 
equivalent. 

Safranin.— 

Safranin. * S* 

95 per cent alcohol. 50 c * c * 

Water. 50c * c * 






















336 


METHODS IN PLANT HISTOLOGY 


Safranin (another formula).— 

Dissolve 1 g. of alcohol-soluble safranin in 100 c.c. of absolute alcohol; 
dissolve 1 g. of water-soluble safranin in 100 c.c. of distilled water. Mix 
equal parts of the two solutions. 

Gentian-Violet.— 

Gentian-violet. 1 g. 

95 per cent alcohol.. 20 c.c. 

Water. 80 c.c. 

Anilin oil. 3 c.c. 

A 1 per cent solution in water keeps better. 

A 1 per cent solution in clove oil is worth a thorough trial. 

Pyoktanin.— -This is sold by E. Merck, in Darmstadt, Germany. 
Dissolve 1 g. of pyoctanin in 30 c.c. of water. 

Orange G.— 

Orange G. 1 g. 

Water. 100 c.c. 

For most purposes a 1 per cent solution in clove oil is preferable. 
It is easier to get a solution by dissolving 1 g. of orange in 100 c.c. 
of absolute alcohol; then add 100 c.c. of clove oil. Let the absolute 
alcohol evaporate until there is left about 100 c.c. of the solution. 

Gold Orange.— 

Gold orange . 1 g. 

Clove oil. 100 c.c. 

Bismarck Brown.—• 

Bismarck brown. 2 g. 

70 per cent alcohol. 100 c.c. 

Nigrosin.— 

Nigrosin . 1 g. 

Water. 100 c.c. 

Gram’s Solution.—■ 

Iodine . 1 g. 

Iodide of potassium. 2 g. 

Water. 300 c.c. 

















FORMULAS FOR REAGENTS 


337 


MISCELLANEOUS 
Mayer’s Albumen Fixative.— 

White of egg (active principle). 50 c.c. 

Glycerin (to keep it from drying up). 50 c.c. 

Salicylate of soda (to keep out bacteria, etc.). 1 g. 

Shake well and filter. 

Land’s Gum Fixative.—• 

Gum arabic. 1 g. 

Potassium bichromate. 1 g. 

Water. 98 c.c. 

Dissolve the gum in water and add the bichromate of potash; 
or dissolve the gum in half the quantity of water and the bichromate 
of potash in the other half, and mix just before using. Le Page’s 
liquid glue may be used instead of the gum arabic. 

Land’s Tropical Developer.— 

Hydrokinon. 8 g. 

Metol. 3 g. 

Sodium Sulphite (dry). 30 g. 

(60 g. if crystals are used) 

Sodium carbonate (dry). 30 g. 

(90 g. if crystals are used) 

Potassium bromide. 2 g. 

Water. 1,000 c.c. 

Land’s Contrast Developer.— 

Hydrokinon. 20 g. 

Sodium sulphite (dry). 00 g. 

Sodium carbonate (dry). 140 g. 

Potassium bromide. 12 g. 

Water. 1,000 c.c. 

Schultze’s Maceration Fluid.— 

The ingredients are nitric acid and potassium chlorate. They are mixed 
only as the reagent is applied. See chapter on “Special Methods” (chap. xi). 

Jeffrey’s Maceration Fluid.—• 

Nitric acid (10 per cent in water). 50 c.c. 

Chromic acid (10 per cent in water). 50 c.c. 

For directions, see chapter xii. 





















338 


METHODS IN PLANT HISTOLOGY 


Fehling’s Solution.— 


. f Cupric sulphate. 3 g. 

\ Water. 100 c.c. 

-n f Sodium potassium tartrate (Rochelle salts).. 16 g. 

\ Water. 100 c.c. 

p j Caustic soda. 12 g. 

\ Water. 100 c.c. 


Keep it in three bottles labeled A, B, and C. When needed for 
use, add 10 c.c. of water to 5 c.c. from each of the three bottles. 


Millon’s Reagent.— 

Mercury. 1 c.c. 

Concentrated nitric acid. 9 c.c. 

Water. 10 c.c. 


Cuprammonia.— 

Prepare by pouring 15 per cent ammonia water upon copper turnings 
or filings. Let it stand in an open bottle. 

Phloroglucin.— 

Use a 5 per cent solution in water or alcohol. 

Celloidin.— 

To make a 2 per cent solution, add one tablet of Schering’s celloidin and 
enough ether-alcohol (equal parts absolute alcohol and ether) to make the 
whole weigh 2,000 g. 

Where only a small quantity is needed, shave off 2 g. of celloidin and add 
100 c.c. of ether alcohol. 

Eycleshymer’s Clearing Fluid.—• 

Mix equal parts of bergamot oil, cedar oil, and carbolic acid. 

Cellulose Acetate.— 


Cellulose acetate. 12 g. 

Pure acetone. 100 c.c. 


Mrs. Williamson recommends the cellulose acetate sold by Cellon, 
Ltd., 22 Cork Street, London, W. I., England. 

Glycerine Jelly.—• 

One part (by weight) of finest French gelatin is left for 2 hours in 6 parts 
(by weight) of distilled water. Add 7 parts of glycerin and for every 100 g. 












FORMULAS FOR REAGENTS 


339 


of the mixture add 1 g. of concentrated carbolic acid. Warm the whole 
mixture for 15 minutes, stirring all the time, until all the flakes produced 
by the carbolic acid have disappeared. Filter, while still warm, through a 
fine-meshed cheese-cloth. 

Venetian Turpentine.— 

To make a 10 per cent solution, add 90 c.c. of absolute alcohol to 10 c.c. 
of thick Venetian turpentine. Stir it with a glass rod. Guess at the amount 
of turpentine, for it is not easy to clean things which have contained Venetian 
turpentine. 

The following need no formulas: acetic acid, hydrochloric acid, 
nitric acid, sulphuric acid, carbolic acid, hydrofluoric acid, acetone, 
chloroform, ether, xylol, cedar oil, clove oil, bergamot oil, turpentine, 
glycerin, paraffin, balsam. 


BIBLIOGRAPHY 

It is not our purpose to give a complete list of literature which might 
be useful, but merely to call attention to some papers and books which will 
extend the student’s knowledge in various directions. Some of the papers 
are cited because they will aid in collecting material and in studying the prepa¬ 
rations rather than for any specific technical methods; for we assume that 
preparations are made to study and not merely for the pleasure of making 
a good mount. 

It will be worth while to watch the reports which are being published in 
Science by the Committee on the Standardization of Biological Stains. The 
reports which have been published as this book goes to press are given below 
under the name of H. J. Conn, the chairman of the Committee. His principal 
collaborators are L. W. Sharp, F. B. Mallory, S. L. Kornhauser, and J. A. 
Ambler. 

Bailey, I. W., Microtechnique for woody structures. Bot. Gaz. 49:57-58. 
1910. 

Belling, John, On counting chromosomes in pollen mother cells. Amer. 
Nat. 55:573-574. 1921. 

Benedict, H. M., An imbedding medium for brittle or woody tissues. 
Bot. Gaz. 52:232! 1911. 

Blackman, V. H., Congo red as a stain for Uredineae. New Phyt. 4:173- 
174. 1905. 

Boyce, J. S., The imbedding and staining of diseased wood. New Phyt. 
8:432-436. 1918. 

Bryan, George S., The archegonium of Sphagnum subsecundum. Bot. Gaz. 
59:40-56. 1915. 

-, The archegonium of Caiharinea angustata. Bot. Gaz. 64:1-20. 1917. 

Bugnon, M. P., Sur une nouvelle m6thode de coloration Elective des 
membranes vegetales lignifiees. C. R. Acad. Sci. Paris 168:62-64. 
1919. 

Church, Margaret, Celloidin paraffin method. Sci. N.S. 47:640. 1918. 
Conn, H. J., An investigation of American stains. Jour. Bact. 7:127-148. 
1922. 

-, Dye solubility in relation to staining solutions. Sci. N.S. 57:638- 

639. 1923. 

-(Chairman) and Commission on Standardization of Biological Stains 

have published the following reports in Science: 

The standardization of biological stains. January 13, 1922. 

340 





BIBLIOGRAPHY 


341 


The production of biological stains in America. 53:289-290. 1921. 

The present supply of biological stains. 56:562-563. 1922. 
Collaborators in the standardization of biological stains. 56:594-596. 
1922. 

Preliminary report on American biological stains.” 56:August 11, 1922. 
American eosins. 56:689-690. 1922. 

The preparation of staining solutions. 57:15-16. 1923. 

Safranin and methyl green. 57:304-305. 1923. 

Dye solubility in relation to staining solutions. 57:41-42. 1923. 
Standardized nomenclature of biological stains. 57:743-746. 1923. 
Certified methylene blue. 58:41-42. 1923. 

Coupin, H., Sur le montage de quelques preparations microscopiques. 
Rev. Gen. Bot. 31:109-114. 1919. 

Cowdry, E. V., Microchemical constituents of protoplasm. Carnegie Inst. 
Wash. Publ. 271. 1918. 

Crocker, E. C., An experimental study of the significance of “lignin” 
color reactions. Jour. Indust, and Eng. Chem. 13:625-627. 1921. 
Davis, W. H., Staining germinating spores. Phytopath. 12:492-494. 1922. 
Degener, Otto, Four new stains of Lycopodium prothallia. Bot. Gaz. 
77:89-95. 1924. 

De Zeeuw, R., The value of double infiltration in botanical microtechnique. 

Papers Mich. Acad. Sci. 1:83-84. 1923. 

Dickson, B. T., The differential staining of plant pathogen and host. Sci. 
N.S. 52:63-64. 1920. 

Dowson, W. T., A new method of paraffin infiltration. Ann. Bot. 36:577- 
578. 1922. 

Durand, E. J., The differential staining of intercellular mycelium. Phyto¬ 
path. 1:129-130. 1911. 

Farmer, J. B., and Shove, Dorothy, On the structure and development of 
the somatic and heterotype chromosomes of Tradescantia virginica. 
Quart. Jour. Mic. Sci. 48:559-569. 1905. 

Gerry, E., and Diemer, M. E., Stains for the mycelium of molds and other 
fungi. Sci. N.S. 54:629-630. 1921. 

Hill, J. B., A method for dehydration of histological material. Bot. Gaz. 
51:255-256. 1916. 

Hubert, E. E., A staining method for hyphae of wood-inhabiting fungi. 
Phytopath. 12:440-441. 1922. 

Karston, G., tJber embryonales Wachstun und seine Tagesperiode. Zeit- 
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Kellicott, W. E., The daily periodicity of cell division and of elongation in 
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342 


METHODS IN PLANT HISTOLOGY 


Kornhauser, S. I., Celloidin paraffin method. Sci. 44:57-58. 1916. 

Land, W. J. G., Microtechnical methods. Bot. Gaz. 59:397-401. 1915. 

-, A method of controlling the temperature of the paraffin block and 

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Langdon, LaDema Mary, Sectioning hard woody tissues. Bot. Gaz. 
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Lee, H. N., The staining of wood fibers for permanent microscopic mounts. 
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Mann, Albert, The preparation of unbroken pollen mother cells and other 
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Metz, C. W., A simple method for handling small objects in making micro¬ 
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Mollendorf, W. yon, Vitale Farbungen an tierischen Zellen. Ergebn. 

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BIBLIOGRAPHY 


343 


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INDEX 













/ 




































INDEX 

[The references are to pages. Italic figures indicate illustrations. 


Abies, 284 
Achlya, 213 
Aecidium, 223, 224 
Albugo, 214 
Alcohol, 19 
Anabaena, 172 
Anilin blue, 62 
Angiopteris, 258, 259 

Angiosperms: stem, 290; root, 292; leaf, 295; 
floral development, 296; Spermatogenesis, 
298, 299; oogenesis, 303, 304; fertilization, 
307, 309; embryo, 309 
Antheridia of Bryophytes, 232, 239 
Anthoceros, 237 
Anthrax, 167 
Araucaria, 284 

Archegonia of Bryophytes, 233 239, 241 

Asclepias, 309 

Ascomycetes, 216 

Aspergillus, 217 

Azolla, 266, 267 

Bacteria, 166 
Basidiomycetes, 221 
Batrachospermum, 205 
Beggiatoa, 168 
Benda’s fluid, 27 
Bensley’s formula, 3-2 
Bibliography, 340 
Boletus, 227 

Botrychium, 256, 258, 259 
Bouin’s fluid, 29 
Bryophytes, 229 

Canaliculi, 14®> 143 
Carmine, 52 
Carnoy’s fluid, 20 

Capsella: floral development, 296, embryo, 310 
Cataloging preparations, 318 
Cellulose acetate, 130 

Ceratozamiaz, 109; pollen tubes, 272; sperm, 274 

Chara, 194, 195 

Chlorophyceae, diagram, 196 

Chondriosomes, 142 

Chromic acid, 21 

Cilia, 141 

Cirsium, floral development, 298 
Cladophora, 191 
Class list of preparations, 320 
Clearing agents, 36 


Clove oil, 38 
Coleochaete, 193 
Collema, 221 

Coniferales, 278; stem, 279, 281; root, 280; 

leaves, 280; oogenesis, 285; embryo, 289 
Coprinus, 225, 227 
Corallina, 208 
Corrosive sublimate, 29 
Covers, 17 
Crucibulum, 227 
Cucurbita, 291 
Cutleria, 201 

Cutting, paraffin, 116, 117 
Cyanin, 60 
Cyathus, 227 
Cycas, 269 

Dehydrating, 33 
Desmids, 183 
Desmotrichum, 200 
Diatoms, 180, 182 
Dicksonia, 81 
Dictyota, 203, 204 
Dioon, 270 
Diospiros, 139 

Ectocarpus, 200 

Endosperm of Angiosperms, 308 

Epidermis, 94 

Equisetum, 251, 252 

Erigeron, floral development, 298 

Erysiphe, 219 

Erysipheae, 218 

Erythronium, 302 

Eurotium, 217 

Eycleshymer’s clearing fluid, 39 

Fertilization, Lilium, 307, 309 
Filicales, 254 

Fixative; Land’s, 119; Mayer’s, 118; Szom- 
bathy’s, 120 

Flemming’s fluid, 26; triple stain, 63 
Formalin, 31 

Formulas for reagents, 327 
Fucus, 202 
Funaria, 240, 241 
Fungi, 209 

Gentian-violet, 59 
Gilson’s fluid, 225 
Ginkgo, 276 


347 


348 


METHODS IN PLANT HISTOLOGY 


Gleichenia, 257 
Gloeotrichia, 170, 171 
Glycerine jelly, 99 
Gnetales, 289 
Gram’s solution, 167 
Gymnosporcmgium, 226 

Haematoxylin, 43 
Hanging drop cultures, 77, 78 
Hepaticae, 229 
Hermann’s fluid, 28 
Holder, for Gillette blades, 9 
Hones, 11 
Hydnum, 227 
Hydrodictyon, 187, 188 
Hydrofluoric acid, 92 

Imbedding: in paraffin, 114, 115; in celloidin, 
125 

Iodine, 31 
Isoetes, 230 

Jeffrey’s maceration method, 138 

Labels, 87, 319 
Laminaria, 200 

Land’s contrast developer, and tropical devel¬ 
oper, 155 

Land’s gelatin method, 137 
Lantern slides, 152 
Lenticels, 291 
Licent’s formula, 26 
Lichens, 221 
Light, artificial, 317 
Light green, 61 
Lignin, 81 
Ligustrum, leaf, 296 

Lilium: spermatogenesis, 298, 299; oogenesis 
304, 305, 306 
Lycoperdon, 227 
Lycopodium, 244, 246 

Magdala red, 58; with anilin blue, 105 

Marchantia, 230, 233, 234, 236 

Marsilia, 266, 267 

Microchemical tests, 78 

Micrometry, 312 

Microscope, 6, 313 

Microsphaera, 218, 219 

Microtome, 7, 108 

Merkel’s fluid, 28 

Mnium, 242 

Mucor, 209 

Musci, 238 

Myxomycetes, 164 

Nemalion, 205, 206 
Nidularia, 227 
Nostoc, 170 
Nummularia, 220 


Oedogonium, 192, 193 
Orange, 62 
Oscillatoria, 169 

Osmunda: stem 257; sporangia, 260; arche- 
gonia; 264, 265 

Paraffin: bath, 13; melting-point, 39 

Parthenogenesis, 311 

Pellia, 232, 236 

Penicillium, 218 

Petrifactions, 133 

Peziza, 216 

Photomicrographs, 145 
Phyllactinia, 219 
Physcia, 221 
Phytelephas, 141 
Picea, 282 
Pilularia, 266 

Pinus, 285, 286; ventral canal cell, 287; 

embryo, 288 
Pistia, root, 295 
Plasmodiophora, 228 
Pleurococcus, 188 
Plumbagella, 308 
Polyporus, 227 
Polysiphonia, 207 
Power’s methods, 178 

Prothallia: of ferns, 261; Costello’s method, 263 

Protonema, 238 

Protoplasmic connections, 139 

Prunus, floral development, 297 

Pteridophytes, 244 

Pteris, 255, 262, 263 

Ptilidium, 231 

Puccinia, 222, 223, 224, 236 

Ranunculus, root, 294 
Rhizopus, 209, 211 
Rhodophyceae, 205 
Riccia, 229, 235 
Rivularia, 170 
Rusts, 222 

Saccharomyces, 215 

Sagittaria, 311 

Salvinia, 266 

Saprolegnia, 212, 213 

Scenedesmus, 186, 187 

Schultze’s maceration method, 137 

Sealing, 97, 98 

Sedum, 94 

Selaginella, 247, 249 

Slides, 17 

Smilax, root, 295 

Smuts, 221 

Sparganium, root, 295 
Spermatophytes, 269 
Sphacelaria, 199 
Sphaerotheca, 218 


INDEX 


349 


Sphagnum, 242, 243 

Sphenophyllum, 136 

Spirogyra, 96, 102, 184, 185 
Sporodinia, 212 

Sporophyte of Bryophytes, 234, 242 
Staining dishes, 15 

Stony tissues, 133 

Syringa, leaf, 296 

Turntable, 14 

Tyloses, 291 

Uncinula, 218, 219 

Vaucheria, 189 

Venetian turpentine, 101 

Vicia Faba, root, 295 

Volvox, 176, 178; Power’s methods, 178, 179 

Taraxacum, 298 

Temporary mounts, 76 

Thermostat, 12 

Thick sections, 135 

Tilia, 291 

Tolypothrix, 169, 170 

Tradescantia, root, 292 

Trichia, 164 

Trillium, root, 292 

Tube length, 313, 314 

Washing, 24 

Xylaria, 220 

Xylol, 36 

Yeast, 215 

Zamia, 269, 276 

Zea Mais, 294 

Zygnema, 184, 185 

Zygorhynchus, 212 


PRINTED IN THE U.S.A. 
























































































